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.

<table><tbody><tr><td>Large bath chamber with slice supports</td><td>Warner Instruments</td><td>RC-27L</td><td>For ex vivo imaging chamber</td></tr><tr><td>Standard Slice Supports</td><td>Warner Instruments</td><td>SS-3</td><td>For ex vivo imaging chamber</td></tr><tr><td>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></tr><tr><td>Suction Tube, Series 20 Classic Design, left handed</td><td>Warner Instruments</td><td>ST-1L</td><td>For ex vivo imaging chamber</td></tr><tr><td>Solution In-line heater/cooler</td><td>Warner Instruments</td><td>SC-20</td><td>To regulate perfusate temperature during imaging</td></tr><tr><td>Bipolar temperature controller</td><td>Warner Instruments</td><td>CL-100</td><td>To regulate perfusate temperature during imaging</td></tr><tr><td>Liquid Cooling System</td><td>Warner Instruments</td><td>LCS-1</td><td>To regulate perfusate temperature during imaging</td></tr><tr><td>Cable assembly for heater controllers</td><td>Warner Instruments</td><td>CC-28</td><td>To regulate perfusate temperature during imaging</td></tr><tr><td>Replacement bead thermisitor for CC-28 cable</td><td>Warner Instruments</td><td>TS-70B</td><td>To regulate perfusate temperature during imaging</td></tr><tr><td>Magnetic holder with suction tubing</td><td>Bioscience Tools</td><td>MTH-S</td><td>To hold the stainless steel vacuum suction tubing&amp;nbsp;</td></tr><tr><td>Adjustable holder</td><td>Bioscience Tools</td><td>MTH</td><td>To hold the temperature probe</td></tr><tr><td>clear silicone sealant</td><td></td><td></td><td>For ex vivo imaging chamber</td></tr><tr><td>superglue</td><td></td><td></td><td>For ex vivo imaging chamber</td></tr><tr><td>thin plexiglass strips</td><td></td><td></td><td>For ex vivo imaging chamber</td></tr><tr><td>nile red</td><td>Life Technologies</td><td>N-1142</td><td>For labeling myelin</td></tr></tbody></table>

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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Research

JoVE Journal

Neuroscience

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

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Intravital Imaging of Axonal Interactions with Microglia and Macrophages in a Mouse Dorsal Column Crush Injury

Traumatic spinal cord injury causes an inflammatory reaction involving blood-derived macrophages and central nervous system (CNS)-resident microglia. Intra-vital two-photon microscopy enables the study of macrophages and microglia in the spinal cord lesion in the living animal. This can be performed in adult animals with a traumatic injury to the dorsal column. Here, we describe methods for distinguishing macrophages from microglia in the CNS using an irradiation bone marrow chimera to obtain animals in which only macrophages or microglia are labeled with a genetically encoded green fluorescent protein. We also describe a injury model that crushes the dorsal column of the spinal cord, thereby producing a simple, easily accessible, rectangular lesion that is easily visualized in an animal through a laminectomy. Furthermore, we will outline procedures to sequentially image the animals at the anatomical site of injury for the study of cellular interactions during the first few days to weeks after injury.

Two-photon intravital imaging can be used to investigate interactions among different cell types in the spinal cord in their native tissue environment in a bone marrow chimeric animal with a dorsal column traumatic spinal cord crush injury.

Traumatic spinal cord injury causes an inflammatory reaction involving blood-derived macrophages and central nervous system (CNS)-resident microglia. ...

Photothrombosis-induced Focal Ischemia as a Model of Spinal Cord Injury in Mice

Spinal cord injury (SCI) is a devastating clinical condition causing permanent changes in sensorimotor and autonomic functions of the spinal cord (SC) below the site of injury. The secondary ischemia that develops following the initial mechanical insult is a serious complication of the SCI and severely impairs the function and viability of surviving neuronal and non-neuronal cells in the SC. In addition, ischemia is also responsible for the growth of lesion during chronic phase of injury and interferes with the cellular repair and healing processes. Thus there is a need to develop a spinal cord ischemia model for studying the mechanisms of ischemia-induced pathology. Focal ischemia induced by photothrombosis (PT) is a minimally invasive and very well established procedure used to investigate the pathology of ischemia-induced cell death in the brain. Here, we describe the use of PT to induce an ischemic lesion in the spinal cord of mice. Following retro-orbital sinus injection of Rose Bengal, the posterior spinal vein and other capillaries on the dorsal surface of SC were irradiated with a green light resulting in the formation of a thrombus and thus ischemia in the affected region. Results from histology and immunochemistry studies show that PT-induced ischemia caused spinal cord infarction, loss of neurons and reactive gliosis. Using this technique a highly reproducible and relatively easy model of SCI in mice can be achieved that would serve the purpose of scientific investigations into the mechanisms of ischemia induced cell death as well as the efficacy of neuroprotective drugs. This model will also allow exploration of the pathological changes that occur following SCI in live mice like axonal degeneration and regeneration, neuronal and astrocytic Ca2+ signaling using two-photon microscopy.

Photothrombosis is a minimally invasive and highly reproducible procedure to induce focal ischemia in the spinal cord and serves as a model of spinal cord injury in mice.

Spinal cord injury (SCI) is a devastating clinical condition causing permanent changes in sensorimotor and autonomic functions of the spinal cord (SC) ...

Medicine

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 Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its incidence is still considerably high and therefore, influences patient outcome. Microcirculation plays a key role in tissue perfusion and oxygen supply and is often dissociated from macrohemodynamics. Thus, direct evaluation of spinal cord microcirculation is essential for the development of microcirculation-targeted therapies and the evaluation of existing approaches in regard to spinal cord microcirculation. However, most of the methods do not provide real-time assessment of spinal cord microcirculation. The aim of this study is to describe a standardized protocol for real-time spinal cord microcirculatory evaluation using laser-Doppler needle probes directly inserted in the spinal cord. We used a porcine model of ischemia/reperfusion to induce deterioration of the spinal cord microcirculation. In addition, a fluorescent microsphere injection technique was used. Initially, animals were anesthetized and mechanically ventilated. Thereafter, laser-Doppler needle probe insertion was performed, followed by the placement of cerebrospinal fluid drainage. A median sternotomy was performed for exposure of the descending aorta to perform aortic cross-clamping. Ischemia/reperfusion was induced by supra-celiac aortic cross-clamping for a total of 48 min, followed by reperfusion and hemodynamic stabilization. Laser-Doppler Flux was performed in parallel with macrohemodynamic evaluation. In addition, automated cerebrospinal fluid drainage was used to maintain a stable cerebrospinal pressure. After completion of the protocol, animals were sacrificed, and the spinal cord was harvested for histopathological and microsphere analysis. The protocol reveals the feasibility of spinal cord microperfusion measurements using laser-Doppler probes and shows a marked decrease during ischemia as well as recovery after reperfusion. Results showed comparable behavior to fluorescent microsphere evaluation. In conclusion, this new protocol might provide a useful large animal model for future studies using real-time spinal cord microperfusion assessment in ischemia/reperfusion conditions.

Spinal cord microcirculation plays a pivotal role in spinal cord injury. Most methods do not allow real-time assessment of spinal cord microcirculation, which is essential for the development of microcirculation-targeted therapies. Here, we propose a protocol using Laser-Doppler-Flow Needle probes in a large animal model of ischemia/reperfusion.

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its ...

Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes ...

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

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Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae