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

The overall goal of the following experiment is to assess acute axonal degeneration mechanisms in real time. This is achieved by isolating thigh one YFP positive neuro and spinal cord tissue, while continually perfusing the tissue with oxygenated low calcium, artificial cerebral spinal fluid. Then the spinal cord tissue is aligned under a two photon microscope objective while maintaining physiological conditions, and the myelin is stained with the lipophilic fluorescent dye Nile red.

Next, a micros size lesion is created on the dorsal surface of a dorsal column by adjusting the laser power in order to transect myelinated axons within the columns. The results show retraction of the damaged axons, peri axonal, myelin swelling, and other cellular responses based on quantitative measurements of the images. So the main advantage of this technique over existing methods is that we use the whole cervical spinal cord and therefore the dorsal column fibers remain intact.

So this avoids damage to the underlying central myelinated fibers, which commonly occurs during white matter strip preparation. Demonstrating this procedure will be Dr.Starlin ada, a postdoctoral fellow in my laboratory who really championed this technique To prepare for the intracardiac perfusion. Make low calcium, artificial cerebral spinal fluid, or A CSF with two times concentrated stock BNC solutions for profusion of the ex vivo spinal cord.

During imaging, prepare normal A CSF with two times stock solutions A and B in a glass bottle. Offer these solutions to pH 7.4. Keep the low calcium A CSF chilled near the dissection area and keep the normal A CSF at room temperature near the microscope.

Bubble both solutions with car and gas for half an hour prior to use and bubble them continuously while they're in use. Next, prepare the open bath perfusion chamber for imaging. First, connect the inline heaters metal output to the chamber.

Then adhere the imaging chamber to the spill chamber with a good amount of tacky material so that there is a potential to slightly adjust the position of the imaging chamber. Secure the inline heaters feedback temperature probe near the fluid inlet and secure the stainless steel suction tube to the vacuum line at the chamber outlet. Now turn on the vacuum pump and the perfusion pump.

Once the chamber fills with A CSF, dial the pump down to 1.5 to two milliliters per minute. Then control the A CF volume in the chamber by the position of the vacuum line. To adjust euthanize six to 10 week old TH one YFP transgenic mouse.

First, confirm there is no reflexive toe pinch response. Then shave the area covering the spinal cord and brain. After shaving, wipe the exposed skin with Betadine and 70%ethanol.

Now spray the chest and abdominal area with 70%ethanol and perfuse the animal trans Cardi with carbogen bubbled low calcium A CSF. When the liver starts to pale, attach the perfusion needle into the heart with a clip. Then turn the animal over and wipe down the dorsal skin side with 70%ethanol.

Make an incision along the midline from the nose to the hip to expose the skull cap and the tissue above the vertebrae. Use pins to pull aside the skin and secure the animal at the nose and haun. Now move the preparation to a dissection microscope there with scissors firmly cut across the dorsal surface of the skull at the level of the olfactory bulbs.

Insert the scissors tip into the newly made incision and to cut along both of the skull cap edges to expose the cerebellum. Do not completely excise the skull cap. Lift up the skull cap exposing the brain and carefully cut the left and right edges of the inter parietal bone over the cerebellum.

Now, while gently pulling the skull cap and dorsal vertebral column surface, make cuts just inferior to the posterior nerve roots to further expose the spinal cord. Then using finely tipped scissors, carefully cut the vertebrae bilaterally. At a 45 degree angle, it is vital to avoid damage to the dorsal columns.

These bright white fibers are easily damaged without being careful, continued gently pulling back the excised bone and muscle while cutting the vertebrae. Cut one to two centimeters to isolate a segment of cervical spinal cord tissue suitable for imaging. Then use scissors to cut the tissue and bone parallel to the vertebral column.

Also clear the tissue underneath the column. Make these cuts as level as possible because it is very important that the tissue lies flat in the chamber level for imaging. Finally, use the number 11 scalpel to transect the brainstem past the termination of the grass ulu fibers and at the upper thoracic level.

Then use scissors to cut the tissue and bone at these two points. The isolated tissue should contain two thirds of the vertebral column encompassing the cordal brainstem, cervical enlargement, and the upper thoracic spinal cord. Transfer the isolated spinal column to the ex vivo imaging chamber and gradually increase the perfuse eight temperature to between 36 and 37 degrees Celsius over the next hour.

Maintain this temperature during imaging. One critical aspect of this protocol is to ensure that you isolate the spinal cord without damaging the dorsal column fibers. The second most critical aspect of this protocol is to ensure that the spinal cord is continuously perfused with oxygenated artificial cerebral spinal fluid.

Using a modified tissue slide holder, carefully secure the vertebral column in the chamber. It is important to avoid touching the exposed spinal cord, dorsal columns, or putting pressure on them with the liker threads threads. Now to stain the myelin, briefly stop the solution flow and remove the vacuum line.

Add five to 10 microliters of five millimolar Nile red dye near the cordal end of the spinal cord. Then mix the solution gently throughout the imaging chamber and allow the tissue to incubate for a minute or two. Once stained, return the flow of oxygenated A CSF to between 1.5 and two milliliters per minute and replace the vacuum line using a two photo microscope.

Image the spinal cord using a water immersion objective. Slowly lower the objective to the A CSF surface, and then bring the spinal cord into focus under AF fluorescent light. Now switch to scanning mode for baseline imaging of the ascending grass.

Oculus myelinated fibers. To excite the YFP and Nile red, set the excitation to 950 nanometers and isolate the fluorescent emissions with DI chronic and bend pass filters. If within an hour of imaging you notice that more than 3%of the axons have steroids, this means that the spinal cord has been damaged during isolation.

It is now best to discard the preparation To injure the tissue with the laser. First, locate an area of the tissue that appears to be level. Then increase the viewing area magnification to 30.3 x.

Tune the laser to 800 nanometers and increase its power to about 110 milliwatts. Now perform five field of view laser scans. The results should be a complete 20 micron diameter ablation.

Check the injury under standard imaging settings with the laser at 17 milliwatts and at 20.08 x magnification. If the axonal fibers at the ablation are not completely transected, discard the preparation. Now, use the time-lapse settings with Z captures to make dynamic injury response recordings.

The preparation can be imaged for up to 24 hours. A spinal cord segment was isolated and positioned as described at baseline T-P-L-S-M. Recording show myelinated grass articular Saxons as parallel aligned YFP positive axons in sheath dinar red stained myelin with few axonal steroids or other morphological signs of axonal degeneration.

Fine details can be seen such as the nose of ROM VA highlighted by arrowheads. 40 minutes later after laser induced spinal cord injury. Most of the transected axons form characteristic axonal end bulbs and retract from the injury site axons adjacent to the injury site undergoes secondary degeneration and axonal swelling became more prominent.

Unlike the quarterly retracting fibers, most of the retracting axonal end bulbs located distally from the lesion and steroids were strongly labeled with Nile Red suggesting an environmental change within the axon. At seven hours, the axons continued retracting away from the lesion site. However, this is more evident in end bulbs located distal or rostral from the lesion than in end bulbs located proximal or cordal to the lesion.

Ultimately, axons that were initially spared gradually underwent delayed secondary degeneration. After watching this video, you should have a good understanding on how to isolate the neuro and spinal cord and perform a laser induced spinal cord injury. Therefore, you'll be able to examine the dynamic aspects of both primary and secondary axonal degeneration in real time.

The information gained from using this model system may lead to important mechanistic understanding of both axon and myelin damage after injury.