DPS acknowledges past and present support in part from grant #2665 and #2934, respectively, from the PVA Research Foundation. PKS is an Alberta Innovates &#8211; Health Solutions Scientist, operating funds were provided by the Leblanc Chair for Spinal Cord Research, University of Calgary.

NOTE: All animal procedures were performed under guidelines set by the Institutional Animal Care and Use committee at the University of Louisville, adhering to Federal regulations.
1. Preparation of Low Ca2+ and 2 mM Ca2+ Artificial Cerebrospinal Fluid (aCSF) Perfusates
<ol>
	<li>Prepare 2x low Ca2+ (0.1 mM) Stock C buffer, 2x normal Ca2+ (2 mM) Stock A buffer, and 2x Stock B buffer as described in Table 1. The individual stock solutions allow modification of ion content (e.g., low or zero Ca2+) and can be stored at 4 &#176;C for up to one month.</li>
	<li>For intracardiac perfusion during dissection, add 100 ml of 2x low Ca2+ Stock C and 100 ml of 2x Stock B into a plastic bottle to make 1x low Ca2+ aCSF. Mix and store overnight at 4 &#176;C or make fresh.</li>
	<li>For perfusion of ex vivo spinal cord during imaging, add 500 ml of 2x normal Ca2+ Stock A and 500 ml of 2x Stock B into a 1 L glass bottle to generate 1x aCSF. Mix and store overnight at room temperature or make fresh.</li>
	<li>Adjust pH of aCSF buffers to 7.4. While keeping the aCSF at room temperature, place the low Ca2+ aCSF on ice and then bubble both buffers with carbogen gas (95% O2; 5% CO2) for 30 min prior to use and continuously during perfusion.</li>
</ol>
2. Preparation of Ex Vivo Imaging Chamber
<ol>
	<li>Wrap a heating blanket (no automatic shut off) around the aCSF bottle and adjust heat to low/ medium setting.</li>
	<li>Insert a dedicated aCSF perfusion line into the bottle connected to a perfusion pump and in-line heater which is controlled by a bipolar temperature controller and feedback temperature probe as shown in Figure 1A.</li>
	<li>Attach a flat microscope stage insert (152 x 102 mm) to the 2-photon excitation/confocal microscope stage fitted with a spill container.</li>
	<li>Place a large open-bath perfusion chamber (W x L x H; 12 x 24 x 8 mm) into the spill container to accommodate and provide continuous flow of fresh oxygenated aCSF to the ex vivo spinal cord (see Figure 1B for a suitable ex vivo imaging chamber design).</li>
	<li>Connect the in-line heater&#8217;s metal output port to the ex vivo imaging chamber using suitable tubing. Place a thin absorbent pad underneath the ex vivo imaging chamber to help maintain the temperature of the buffer/tissue during imaging.</li>
	<li>Adhere the ex vivo imaging chamber to the bottom of the spill container/stage insert using removable sticky tack. The malleability of the sticky tack will allow the chamber to be adjusted as needed to ensure the objective&#8217;s light path is perpendicular to the tissue surface. If necessary, use a bullseye level to adjust the chamber.</li>
	<li>Secure an in-line heater feedback temperature probe near the chamber fluid inlet and a stainless steel suction tube connected to a vacuum line at the chamber outlet. Magnetic tape and appropriate micropipette holders are useful in this regard. Turn on the vacuum source.</li>
	<li>Turn on the perfusion pump to start filling the imaging chamber with oxygenated aCSF. Set the perfusion pump to 1.5-2 ml per min.</li>
	<li>Control the volume of aCSF in the imaging chamber by adjusting the vacuum line. It is important that there be enough aCSF in the imaging chamber to cover the spinal cord segment and allow sufficient working distance between a water dipping objective and the tissue surface. Myelin and axons can be detrimentally affected if the tissue segment is not completely submersed in fresh oxygenated aCSF leading to experimenter-induced artifacts.</li>
	<li>Adjust the in-line heater temperature controller so that perfusate temperature in the imaging chamber is near room temperature.</li>
</ol>
3. Dissection of Adult Murine Cervical Spinal Cord
<ol>
	<li>Euthanize an adult 6-10 week old thy1-YFP transgenic mouse (use appropriate strain that has fluorescent protein expression in dorsal root ganglion neurons) by injecting an overdose of the anesthesia sodium pentobarbital (200 mg/kg) via intraperitoneal injection.</li>
	<li>Once under the proper level of anesthesia (e.g., no reaction to toe pinch and loss of blink reflex), carefully shave the skin overlying the spinal cord and brain with surgical clippers. Cleanse skin using betadine and 70% ethanol pads.</li>
	<li>Spray chest/abdomen area with 70% ethanol, then perfuse the subject transcardially using chilled, carbogen-bubbled low Ca2+ aCSF. (See Figure 1C for suggested dissection bench top set up).</li>
	<li>As the liver pales, secure the perfusion needle into the heart using a small clip. Turn the subject over to access the shaved dorsal side of the subject and wipe the shaved area with 70% ethanol.</li>
	<li>Make a dorsal skin incision along the midline starting from the nose to hip using dissection scissors to expose the brain and vertebral column (Figure 2A). Secure the preparation by placing pins at the nose and haunch.</li>
	<li>Using a dissection microscope from this point on, firmly cut across the dorsal surface of the skull at the level of the olfactory bulbs. Insert one of the scissor tips into the lateral edges of the incision and cut along the edges of the skullcap bilaterally until the cerebellum. 
	NOTE: Do not completely excise the skullcap as it is needed later to lift the overlying tissue from the spinal cord throughout the dissection (see Figure 2B).</li>
	<li>Lift the skullcap to expose the brain. Gently cut the left and right edges of the interparietal bone (located at the cerebellum) and pull it back caudally to expose the brainstem/spinal cord (Figure 2C).</li>
	<li>Use fine tipped scissors held at a 45&#176; angle to the table surface and cut the vertebrae bilaterally just inferior to the posterior nerve roots. 
	NOTE: Avoid contact with the bright white dorsal columns. These fibers are the ones that will be imaged (Figure 2D) and can be easily damaged.</li>
	<li>Lift the skullcap and dorsal surface of the vertebral column with fine tipped forceps, gently pulling the overlying tissue in one piece as cuts are made to expose the spinal cord. Continue to cut vertebrae to the upper thoracic level to isolate a 1-2 cm segment of the cervical cord for subsequent imaging (Figure 2E).</li>
	<li>Use scissors to cut the tissue/bone parallel to the vertebral column and clear tissue underneath the column. Make these cuts as level as possible. It is very important that the tissue lies flat in the chamber so that the cord is level for imaging.</li>
	<li>Use a #11 scalpel to transect the brain stem (past the termination of the gracile fasciculus fibers) and at the upper thoracic level to isolate the cervical spinal cord segment (See Figure 2F). Use scissors to cut tissue/bone at these same two points. The isolated tissue should contain 2/3 of the vertebral column encompassing the caudal brainstem, cervical enlargement, and upper thoracic spinal cord (Figure 2G).</li>
	<li>Place the isolated spinal column into a petri dish of carbogen-bubbled, chilled low Ca2+ aCSF. If needed, trim the tissue underneath the spinal column until it lies flat in the dish.</li>
	<li>Transfer the isolated spinal column to the ex vivo imaging chamber and gradually increase the perfusate temperature over 1 hr to 36 - 37 &#176;C. Maintain this temperature during imaging.</li>
</ol>
4. Placement of Ex Vivo Spinal Cord into Imaging Chamber and Myelin Labeling with Nile Red
<ol>
	<li>Carefully secure the vertebral column in the imaging chamber with an appropriate fitting slice hold down modified with a vertical grid platform consisting of fine Lycra threads (remove middle threads of grid platform to allow space for the spinal cord; see Figure 3A).</li>
	<li>Thaw an aliquot of Nile Red (5 mM stock in DMSO and 0.22 &#956;m filtered; can be stored for 1 month at 4 &#176;C, or 6 months at -20 &#176;C in the dark). Just before adding Nile Red to the imaging chamber, reduce the flow of the perfusion pump and adjust the vacuum line to ensure the fluid in the chamber always covers the spinal cord.</li>
	<li>Add 5-10 &#181;l Nile Red stock solution to the imaging chamber near the caudal end of the cord (Figure 3B). Use a 1 ml pipette to gently mix Nile Red throughout the chamber. Allow Nile Red to stay in chamber for 1-2 min, then place vacuum line back and adjust perfusion pump to 1.5-2 ml per min to continuously perfuse the spinal cord.</li>
	<li>Carefully raise the microscope stage and align the objective with the center of the spinal cord segment and medially over the dorsal columns until the water immersion objective is approximately 10 mm from the surface of the aCSF in the chamber. Use the microscope nosepiece focus wheel to slowly bring the objective down until it gently touches the surface of the aCSF. Use an epifluorescent light source to focus the dorsal columns into the field of view (Figure 3C).</li>
</ol>
5. Ex Vivo Imaging of the Mouse Spinal Cord with TPLSM and Laser-induced Spinal Cord Injury (LiSCI)
NOTE: The posterior vein provides a useful marker to center the tissue as the gracile fasciculus fibers (originate from cell body of dorsal root ganglia and ascend from T6 and below along the midline of spinal cord) run parallel to this blood vessel. The thicker YFP+ cervical dorsal root projections also provide a useful marker as the fibers ascend and descend laterally to the YFP+ gracile fasciculus fibers that are smaller in diameter.
<ol>
	<li>Once the spinal cord tissue is aligned appropriately, switch to laser scanning mode to perform baseline imaging of the ascending gracile fasciculus myelinated fibers. To excite both YFP and Nile Red, use a two-photon laser source tuned to 950 nm wavelength (~17 mW measured at the exit of a 25X, 1.1 NA objective) and appropriate dichroics (DM) and bandpass filters to isolate the fluorescent emission of the fluorophores (we use 525/50 DM560, 600/60 DM640, and 685/70, to separate YFP, and Nile Red into yellow-orange, and extended red channels, respectively).
	<ol>
		<li>If more than 3% of the axons show axonal spheroids during baseline imaging (30-60 min), discard the preparation due to experimenter-induced artifacts.</li>
	</ol>
	</li>
	<li>Induce a LiSCI by magnifying the field of view to 30.3X (to create a ~20 &#956;m diameter ablation). Tune the laser to 800 nm, and increase the power of the laser to ~110 mW at the sample. Allow 5 complete field of view laser scans to ensure fibers are completely transected.</li>
	<li>Confirm LiSCI by changing laser settings back to imaging settings (950 nm, 17 mW at the sample, 2.08X) and scan the tissue to visualize the ablation. Verify the diameter of the ablation and that the primary ablated axons are completely transected (see Figure 3D). Use time-lapse settings on the microscope combined with z capture to record the dynamic response of axons and myelin to injury up to 8-10+ hr after injury. 
	NOTE: If the ablation is incomplete (i.e., incomplete transection of fibers), discard the preparation, as determination of primary and secondary injury mechanisms will be confounded. In addition, isolated spinal cord segments can be imaged up to 24 hr post-LiSCI with only mild to moderate LiSCI-independent tissue damage.</li>
</ol>

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K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+axonal+injury:+internodal+nanocomplexes+and+calcium+deregulation.">Mechanisms of axonal injury: internodal nanocomplexes and calcium deregulation.</a> Trends Mol Med. 16, 160-170 (2010).</li><li>Wang, H., Fu, Y., Zickmund, P., Shi, R., Cheng, J. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coherent+anti-stokes+Raman+scattering+imaging+of+axonal+myelin+in+live+spinal+tissues.">Coherent anti-stokes Raman scattering imaging of axonal myelin in live spinal tissues.</a> Biophys J. 89, 581-591 (2005).</li><li>Beirowski, B., Nogradi, A., Babetto, E., Garcia-Alias, G., Coleman, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+axonal+spheroid+formation+in+central+nervous+system+Wallerian+degeneration.">Mechanisms of axonal spheroid formation in central nervous system Wallerian degeneration.</a> J Neuropathol Exp Neurol. 69, 455-472 (2010).</li><li>Shi, R., Asano, T., Vining, N. C., Blight, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+membrane+sealing+in+injured+mammalian+spinal+cord+axons.">Control of membrane sealing in injured mammalian spinal cord axons.</a> J Neurophysiol. 84, 1763-1769 (2000).</li><li>Stirling, D. P., Cummins, K., Wayne Chen, ., R, S., Stys, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axoplasmic+reticulum+Ca(2+)+release+causes+secondary+degeneration+of+spinal+axons.">Axoplasmic reticulum Ca(2+) release causes secondary degeneration of spinal axons.</a> Ann Neurol. 75, 220-229 (2014).</li></ol>

The authors have nothing to disclose.

We describe a method of imaging ex vivo spinal cord myelinated axons (i.e., gracile fasciculus) combined with a laser-induced spinal cord injury to study the dynamic progression of both primary and secondary myelinated axonal degeneration over time. Ex vivo imaging of the surface of the spinal cord overcomes many of the complications associated with in vivo imaging such as motion artifacts and the potential of experimenter-induced hypoxia during prolonged imaging sessions. This protoco...

Degeneration of axons is a prominent cause of morbidity spanning several neurological conditions including neurotrauma, stroke, autoimmune, and neurodegenerative diseases. Unlike the peripheral nervous system (PNS), central nervous system (CNS) axons have a limited capacity to regenerate once injured due to both intrinsic and extrinsic barriers (i.e., inhibitory molecules to axonal growth produced during scar formation and liberated during myelin degeneration)1-7. Although several of these barriers have been extensively explored, therapeutic interventions aimed to prevent CNS axonal degeneration, promote robust axonal regeneration, and restore functional connectively, remain limited.
Axons once separated from their soma undergo a stereotypical process of degeneration known as Wallerian degeneration that is characterized by axonal swelling, spheroid formation and eventual fragmentation (reviewed in 8). In contrast, the proximal stump that remains in continuity with the soma of a transected peripheral axon, forms a swelling at its end, dies back to the nearest node of Ranvier, and can then initiate growth cone formation, a vital prerequisite necessary for subsequent axonal regeneration9-11. In contrast, the proximal axonal endings of many central axons form characteristic &#8220;endbulbs&#8221; or retraction bulbs, fail to form growth cones, and instead retract away from the injury site where they remain for months after injury12-15. In addition to the primary axonal injury, additional axonal damage/loss may also occur to axons that were largely spared from the initial injury. This delayed axonal loss of initially spared axons is referred to as secondary axonal degeneration. This inherent response of CNS axons to injury renders functional axonal regeneration an even more difficult goal to achieve in the brain and spinal cord.
Although hallmarks of axonal injury (e.g., spheroid formation, retraction bulbs) have been well characterized from post-mortem tissue and experimental models of axonal degeneration, elucidation of the molecular mechanisms underlying these dynamic processes has been restricted. Most of these studies relied on static endpoint observations that inherently failed to capture individual axonal responses over time. Though exogenously applied axonal tracers have been useful to elucidate axonal responses from static sections and during live imaging, the availability of genetically encoded axonal markers has greatly improved our ability to visualize axons in real-time using fluorescence microscopy. Indeed, a seminal report from Kerschensteiner and colleagues first provided direct evidence of axonal degeneration and regeneration in vivo using Thy1-GFP-S mice that encode green fluorescent protein in subsets of neurons that send their projections in the dorsal columns of the spinal cord16. Live imaging approaches using two photon laser scanning microscopy (TPLSM) and genetic fluorescent protein labeling of cells of interest continues to provide direct evidence and mechanistic insight into many diverse dynamic processes such as axonal degeneration, Ca2+ signaling, axonal regeneration, astrocyte physiology, microglial physiology, and response to injury17-25.
In contrast to axons, very little is known of myelin responses to injury in real-time. Myelin is a vital component of white matter produced and maintained by oligodendrocytes in the CNS and Schwann cells in the PNS. Myelin insulates 99% of the surface of axons and by doing so provides a high-resistance, low-capacitance protective covering that supports rapid and efficient saltatory impulse propagation, recently reviewed by Buttermore et al.26. To capture the dynamic response of myelin to injury we use the solvatochromic, lipophilic fluorescent dye Nile Red27. The solvatochromic properties of this vital stain allow spectral shifts of its emission spectrum that is dependent on the physico-chemical environment28,29. These properties are useful to gain insight into mechanisms of axomyelinic injury and can be visualized using appropriately selected dichroics and emission filters or resolved using spectral microscopy27. For example, Nile Red&#8217;s emission spectrum is blue-shifted in less polar, lipid-rich environments such as those found in adipocytes and normal CNS myelin (peak emission ~ 580-590 nm)27. In contrast, this vital dye&#8217;s emission spectrum peaks at ~ 625 nm in endbulbs formed as axons undergo axonal dieback27. Although the precise mechanisms underlying these spectral shifts specifically in endbulbs versus normal myelin remain unclear, such spectral changes may reveal underlying alterations in protein accumulation or disorganization leading to exposure of hydrophobic binding sites 27.
While in vivo imaging is the ultimate metric for observing spinal cord axonal injury dynamics in their native environment, it is technically challenging and requires substantial surgical expertise, and often repeat surgeries to expose the dorsal column that may introduce experimental artifacts (e.g., inflammation and scar formation). In addition, costly equipment is often needed to allow suspension and positioning of an intact animal under the microscope objective lens. The animals need to be carefully monitored as well to ensure they remain warm, to ensure fluids are replenished, and to ensure there are no signs of hypoxia due to prolonged anesthetized imaging sessions. The latter is extremely important as axons and myelin absolutely require constant perfusion and adequate oxygen levels to remain viable. However, this is often not reported or monitored in most in vivo studies to date. In addition, movement artifacts due to heart rate and breathing (isoflurane anesthetized adult mouse: ~ 300-450 beats per min (BPM) is optimal to maintain 97-98% oxygen saturation (normal rate ~ 632 BPM) and ~ 55-65 breaths per min (normal rate is ~ 163 breaths per min), respectively))30 encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution fluorescence microscopy challenging as even the fastest laser scans unavoidably are subject to these movement artifacts. Advances in ultrafast resonant scanners combined with an implantable rigid vertebral framed window may allow imaging of the murine spinal cord in awake animals, but faster scan times inevitably reduce the signal to noise ratio degrading image quality. Further improvements in spinal cord imaging techniques as currently used for brain imaging may overcome many of these obstacles and limit potential confounds introduced by inadequate tissue perfusion, e.g., 31-33.
Much of what is known about white matter physiology and mechanisms of white matter injury has been determined using in vitro or ex vivo preparations of white matter from optic nerve, peripheral nerve, and strips of spinal cord white matter34-41. These preparations continue to advance our knowledge of white matter injury mechanisms as they allow controlled changes in environmental factors, controlled application of drugs and reagents, functional assessments using electrophysiology, and direct fluorescence microscopy observations of axons and myelin in living tissue. Yet, some previous approaches to observe axons from spinal cord dorsal column strips or ventral white matter strips unavoidably injure surface axons during the removal stage that may influence the response of closely opposed axons. To capitalize on the experimental manipulations above and avoid damage to the very fibers under investigation, we use an ex vivo cervical spinal cord model as it prevents direct contact of the dorsal aspect of the cord. Thus, the architecture of the pia mater and adjacent superficial dorsal column axons remain viable and unperturbed during isolation.
Here we describe a relatively simple approach that allows direct visualization of central myelinated axons as they dynamically respond to a focal injury in real-time up to 8 - 10+ hr after injury. The laser-induced spinal cord injury (LiSCI) model allows differentiation between primary and secondary axonal injury mechanisms as the primary lesion (ablation site) remains spatially constrained over time. The open-bath imaging chamber is accessible to therapeutic intervention, reagent delivery, and environmental manipulations. Putative axomyelinic protective agents can be quickly assessed in real-time by direct observations versus lengthy and costly experiments involving tissue processing, sectioning, immunostaining, image capture, and analysis and therefore provides a useful surrogate model to assess acute responses and protective manipulations before testing the experimental agents in live animals.

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			<td style="width:341px;">Comments/Description</td>
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			<td height="21" style="height:21px;">Large bath chamber with slice supports</td>
			<td>Warner Instruments</td>
			<td>RC-27L</td>
			<td>For ex vivo imaging chamber</td>
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			<td>Warner Instruments</td>
			<td>SS-3</td>
			<td>For ex vivo imaging chamber</td>
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			<td height="21" style="height:21px;">Plastic Slice hold-down for RC-27L and RC-29 chambers</td>
			<td>Warner Instruments</td>
			<td>SHD-27LP/10</td>
			<td>For ex vivo imaging chamber</td>
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			<td height="21" style="height:21px;width:376px;">Suction Tube, Series 20 Classic Design, left handed</td>
			<td style="width:139px;">Warner Instruments</td>
			<td style="width:113px;">ST-1L</td>
			<td>For ex vivo imaging chamber</td>
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			<td height="21" style="height:21px;">Solution In-line heater/cooler</td>
			<td>Warner Instruments</td>
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			<td>To regulate perfusate temperature during imaging</td>
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			<td style="width:139px;">Warner Instruments</td>
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			<td>To regulate perfusate temperature during imaging</td>
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			<td style="width:139px;">Warner Instruments</td>
			<td style="width:113px;">LCS-1</td>
			<td>To regulate perfusate temperature during imaging</td>
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			<td height="21" style="height:21px;width:376px;">Cable assembly for heater controllers</td>
			<td style="width:139px;">Warner Instruments</td>
			<td style="width:113px;">CC-28</td>
			<td>To regulate perfusate temperature during imaging</td>
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			<td height="21" style="height:21px;width:376px;">Replacement bead thermisitor for CC-28 cable</td>
			<td style="width:139px;">Warner Instruments</td>
			<td style="width:113px;">TS-70B</td>
			<td>To regulate perfusate temperature during imaging</td>
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			<td height="21" style="height:21px;width:376px;">Magnetic holder with suction tubing</td>
			<td style="width:139px;">Bioscience Tools</td>
			<td style="width:113px;">MTH-S</td>
			<td>To hold the stainless steel vacuum suction tubing&#160;</td>
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			<td height="21" style="height:21px;width:376px;">Adjustable holder</td>
			<td style="width:139px;">Bioscience Tools</td>
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			<td>To hold the temperature probe</td>
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			<td height="21" style="height:21px;width:376px;">clear silicone sealant</td>
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			<td height="21" style="height:21px;width:376px;">thin plexiglass strips</td>
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			<td height="21" style="height:21px;width:376px;">nile red</td>
			<td style="width:139px;">Life Technologies</td>
			<td>N-1142</td>
			<td>For labeling myelin</td>
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an ex vivo laser-induced spinal cord injury model to assess mechanisms of axonal degeneration in real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

Details of an appropriate laboratory set up needed to isolate, maintain viability, and image the ex vivo spinal cord is shown in Figure 1. The microscope needs to be equipped with a tunable pulsed femtosecond laser, appropriate dicroics and emission filters, and a water-dipping objective lens with a high numerical aperture (&#8805;1.0). To ensure viability of the spinal cord during the dissection, the procedure should be performed in the presence of chilled oxygenated low Ca2+ aCSF th...

Watch this Scientific Journal Video about An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time at JoVE.com

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

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10.8K Views.  University of Louisville. We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability. 

Video: An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Lateral Fluid Percussion: Model of Traumatic Brain Injury in Mice

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity and mortality. Based on the nature of primary injury following TBI, complex and heterogeneous secondary consequences result, which are followed by regenerative processes 1,2. Primary injury can be induced by a direct contusion to the brain from skull fracture or from shearing and stretching of tissue causing displacement of brain due to movement 3,4. The resulting hematomas and lacerations cause a vascular response 3,5, and the morphological and functional damage of the white matter leads to diffuse axonal injury 6-8. Additional secondary changes commonly seen in the brain are edema and increased intracranial pressure 9. Following TBI there are microscopic alterations in biochemical and physiological pathways involving the release of excitotoxic neurotransmitters, immune mediators and oxygen radicals 10-12, which ultimately result in long-term neurological disabilities 13,14. Thus choosing appropriate animal models of TBI that present similar cellular and molecular events in human and rodent TBI is critical for studying the mechanisms underlying injury and repair.
Various experimental models of TBI have been developed to reproduce aspects of TBI observed in humans, among them three specific models are widely adapted for rodents: fluid percussion, cortical impact and weight drop/impact acceleration 1. The fluid percussion device produces an injury through a craniectomy by applying a brief fluid pressure pulse on to the intact dura. The pulse is created by a pendulum striking the piston of a reservoir of fluid. The percussion produces brief displacement and deformation of neural tissue 1,15. Conversely, cortical impact injury delivers mechanical energy to the intact dura via a rigid impactor under pneumatic pressure 16,17. The weight drop/impact model is characterized by the fall of a rod with a specific mass on the closed skull 18. Among the TBI models, LFP is the most established and commonly used model to evaluate mixed focal and diffuse brain injury 19. It is reproducible and is standardized to allow for the manipulation of injury parameters. LFP recapitulates injuries observed in humans, thus rendering it clinically relevant, and allows for exploration of novel therapeutics for clinical translation 20.
We describe the detailed protocol to perform LFP procedure in mice. The injury inflicted is mild to moderate, with brain regions such as cortex, hippocampus and corpus callosum being most vulnerable. Hippocampal and motor learning tasks are explored following LFP.

Lateral fluid percussion (LFP), an established model of traumatic brain injury in mice, is demonstrated. LFP fulfills three major criteria for animal models: validity, reliability and clinical relevance. The procedure, consisting of surgical craniotomy, fixation of hub followed by induction of injury, resulting in focal and diffuse injuries, is described.

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity and ...

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

M&#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after severe damage. Investigating the regeneration pathways in zebrafish might bring new insights to develop innovative strategies for the treatment of retinal degenerative diseases in mammals. Herein, we focused on the induction of a focal lesion to the outer retina in adult zebrafish by means of a 532 nm diode laser. A localized injury allows investigating biological processes that take place during retinal degeneration and regeneration directly at the area of damage. Using non-invasive optical coherence tomography (OCT), we were able to define the location of the damaged area and monitor subsequent regeneration in vivo. Indeed, OCT imaging produces high-resolution, cross-sectional images of the zebrafish retina, providing information which was previously only available with histological analyses. In order to confirm the data from real-time OCT, histological sections were performed and regenerative response after the induction of the retinal injury was investigated by immunohistochemistry.

M&amp;#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

The zebrafish is a popular animal model to study mechanisms of retinal degeneration/regeneration in vertebrates. This protocol describes a method to induce localized injury disrupting the outer retina with minimal damage to the inner retina. Subsequently, we monitor in vivo the retinal morphology and the M&#252;ller glia response throughout retinal regeneration.

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

An In Vivo Duo-color Method for Imaging Vascular Dynamics Following Contusive Spinal Cord Injury

Spinal cord injury (SCI) causes significant vascular disruption at the site of injury. Vascular pathology occurs immediately after SCI and continues throughout the acute injury phase. In fact, endothelial cells appear to be the first to die after a contusive SCI. The early vascular events, including increased permeability of the blood-spinal cord barrier (BSCB), induce vasogenic edema and contribute to detrimental secondary injury events caused by complex injury mechanisms. Targeting the vascular disruption, therefore, could be a key strategy to reduce secondary injury cascades that contribute to histological and functional impairments after SCI. Previous studies were mostly performed on postmortem samples and were unable to capture the dynamic changes of the vascular network. In this study, we have developed an in vivo duo-color two-photon imaging method to monitor acute vascular dynamic changes following contusive SCI. This approach allows detecting blood flow, vessel diameter, and other vascular pathologies at various sites of the same rat pre- and post-injury. Overall, this method provides an excellent venue for investigating vascular dynamics.

An In Vivo Duo-color Method for Imaging Vascular Dynamics Following Contusive Spinal Cord Injury

We introduce an in vivo imaging method using two different fluorescent dyes to track dynamic spinal vascular changes following a contusive spinal cord injury in adult Sprague-Dawley rats.

Spinal cord injury (SCI) causes significant vascular disruption at the site of injury. Vascular pathology occurs immediately after SCI and continues ...

A Neurosphere Assay to Evaluate Endogenous Neural Stem Cell Activation in a Mouse Model of Minimal Spinal Cord Injury

Neural stem cells (NSCs) in the adult mammalian spinal cord are a relatively mitotically quiescent population of periventricular cells that can be studied in vitro using the neurosphere assay. This colony-forming assay is a powerful tool to study the response of NSCs to exogenous factors in a dish; however, this can also be used to study the effect of in vivo manipulations with the proper understanding of the strengths and limitations of the assay. One manipulation of the clinical interest is the effect of injury on endogenous NSC activation. Current models of spinal cord injury provide a challenge to study this as the severity of common contusion, compression, and transection models cause the destruction of the NSC niche at the site of the injury where the stem cells reside. Here, we describe a minimal injury model that causes localized damage at the superficial dorsolateral surface of the lower thoracic level (T7/8) of the adult mouse spinal cord. This injury model spares the central canal at the level of injury and permits analysis of the NSCs that reside at the level of the lesion at various time points following injury. Here, we show how the neurosphere assay can be utilized to study the activation of the two distinct, lineally-related, populations of NSCs that reside in the spinal cord periventricular region - primitive and definitive NSCs (pNSCs and dNSCs, respectively). We demonstrate how to isolate and culture these NSCs from the periventricular region at the level of injury and the white matter injury site. Our post-surgical spinal cord dissections show increased numbers of pNSC and dNSC-derived neurospheres from the periventricular region of injured cords compared to controls, speaking to their activation via injury. Furthermore, following injury, dNSC-derived neurospheres can be isolated from the injury site &#8212; demonstrating the ability of NSCs to migrate from their periventricular niche to sites of injury.

Here, we demonstrate the performance of a minimal spinal cord injury model in an adult mouse that spares the central canal niche housing endogenous neural stem cells (NSCs). We show how the neurosphere assay can be used to quantify activation and migration of definitive and primitive NSCs following injury.

Neural stem cells (NSCs) in the adult mammalian spinal cord are a relatively mitotically quiescent population of periventricular cells that can be studied ...

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) ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Induction of Complete Transection-Type Spinal Cord Injury in Mice

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, such as cell transplantation methods, are being researched to overcome the debilitating disabilities arising from SCI. Most pre-clinical animal trials are conducted in rodent models of SCI. While rat models of SCI have been widely used, mouse models have received less attention, even though mouse models can have significant advantages over rat models. The small size of mice equates to lower animal maintenance costs than for rats, and the availability of numerous transgenic mouse models is advantageous for many types of studies. Inducing repeatable and precise injury in the animals is the primary challenge for SCI research, which in small rodents requires high-precision surgery. The transection-type injury model has been a commonly used injury model over the last decade for transplantation-based therapeutic research, however a standardized method for inducing a complete transection-type injury in mice does not exist. We have developed a surgical protocol for inducing a complete transection type injury in C57BL/6 mice at thoracic vertebral level 10 (T10). The procedure uses a small tip drill instead of rongeurs to precisely remove the lamina, after which a thin blade with rounded cutting edge is used to induce the spinal cord transection. This method leads to reproducible transection-type injury in small rodents with minimal collateral muscle and bone damage and therefore minimizes confounding factors, specifically where behavioral functional outcomes are analyzed.

This protocol describes how to create a precise laminectomy for induction of stable transection-type spinal cord injury in the mouse model, with minimal collateral damage for spinal cord injury research.

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, ...