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

Summary

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.

Abstract

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.

Introduction

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 regeneration^9-11. In contrast, the proximal axonal endings of many central axons form characteristic “endbulbs” or retraction bulbs, fail to form growth cones, and instead retract away from the injury site where they remain for months after injury^12-15. 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 injury^17-25.

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 environment^28,29. 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’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’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 matter^34-41. 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.

Protocol

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

1. 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 °C for up to one month.
2. 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 °C or make fresh.
3. 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.
4. 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.

2. Preparation of Ex Vivo Imaging Chamber

1. Wrap a heating blanket (no automatic shut off) around the aCSF bottle and adjust heat to low/ medium setting.
2. 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.
3. Attach a flat microscope stage insert (152 x 102 mm) to the 2-photon excitation/confocal microscope stage fitted with a spill container.
4. 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).
5. Connect the in-line heater’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.
6. 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’s light path is perpendicular to the tissue surface. If necessary, use a bullseye level to adjust the chamber.
7. 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.
8. 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.
9. 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.
10. Adjust the in-line heater temperature controller so that perfusate temperature in the imaging chamber is near room temperature.

3. Dissection of Adult Murine Cervical Spinal Cord

1. 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.
2. 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.
3. 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).
4. 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.
5. 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.
6. 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).
7. 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).
8. Use fine tipped scissors held at a 45° 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.
9. 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).
10. 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.
11. 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).
12. 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.
13. Transfer the isolated spinal column to the ex vivo imaging chamber and gradually increase the perfusate temperature over 1 hr to 36 - 37 °C. Maintain this temperature during imaging.

4. Placement of Ex Vivo Spinal Cord into Imaging Chamber and Myelin Labeling with Nile Red

1. 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).
2. Thaw an aliquot of Nile Red (5 mM stock in DMSO and 0.22 μm filtered; can be stored for 1 month at 4 °C, or 6 months at -20 °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.
3. Add 5-10 µl 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.
4. 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).

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.

1. 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).
   1. If more than 3% of the axons show axonal spheroids during baseline imaging (30-60 min), discard the preparation due to experimenter-induced artifacts.
2. Induce a LiSCI by magnifying the field of view to 30.3X (to create a ~20 μm 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.
3. 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.

Results

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 (≥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 that all...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Large bath chamber with slice supports	Warner Instruments	RC-27L	For ex vivo imaging chamber
Standard Slice Supports	Warner Instruments	SS-3	For ex vivo imaging chamber
Plastic Slice hold-down for RC-27L and RC-29 chambers	Warner Instruments	SHD-27LP/10	For ex vivo imaging chamber
Suction Tube, Series 20 Classic Design, left handed	Warner Instruments	ST-1L	For ex vivo imaging chamber
Solution In-line heater/cooler	Warner Instruments	SC-20	To regulate perfusate temperature during imaging
Bipolar temperature controller	Warner Instruments	CL-100	To regulate perfusate temperature during imaging
Liquid Cooling System	Warner Instruments	LCS-1	To regulate perfusate temperature during imaging
Cable assembly for heater controllers	Warner Instruments	CC-28	To regulate perfusate temperature during imaging
Replacement bead thermisitor for CC-28 cable	Warner Instruments	TS-70B	To regulate perfusate temperature during imaging
Magnetic holder with suction tubing	Bioscience Tools	MTH-S	To hold the stainless steel vacuum suction tubing
Adjustable holder	Bioscience Tools	MTH	To hold the temperature probe
clear silicone sealant			For ex vivo imaging chamber
superglue			For ex vivo imaging chamber
thin plexiglass strips			For ex vivo imaging chamber
nile red	Life Technologies	N-1142	For labeling myelin