The goal of this procedure is to perform stable in vivo imaging in the mouse spinal cord using two photon microscopy. This is accomplished by first anesthetizing the animal with an anesthetic mix that achieve a steady yet calm breathing rhythm. The second step of the procedure is to expose the spinal column at the desired level and then perform a laminectomy to expose the underlying spinal cord so that it can be imaged in vivo.
The third step is to place the animal on the spinal stabilization device. The final step is the isolation of the exposed spinal cord segment. In preparation for in vivo imaging, ultimately, results can be obtained that detail the dynamic behavior of cells and their interactions with other cell types or structures in the living spinal cord through time-lapse in vivo imaging using two photon microscopy.
The main advantage of this technique over previous attempts to image the spinal cord in vivo is that it generates road in vivo imaging data that require no post-processing and can be immediately evaluated and used for further analysis. The implications of this technique extend toward treatment following spinal cord injury or disease because, for example, it allows us to study the progression of axonal degeneration or regeneration after, for example, an injury in real time besides spinal cord injury. This technique can be used to to study the progression of inflammatory or neurodegenerative processes in animal models of disease, such as multiple sclerosis or amyotrophic lateral sclerosis, where the disease pathology is prominent in the spinal cord.
Begin the procedure by weighing the animal. Next, anesthetize the animal by injecting the anesthetic mix intraperitoneal. Perform regular toe pinching to ensure deep anesthesia of the animal throughout the experiment and supplement with half the original dose of anesthetic, hourly, or as needed.
Then apply the artificial tears ointment over the eyes to prevent corneal dehydration and damage. Trim the hair on the back of the animal with a trimming device and shave the remaining hair with a sharp blade. After disinfecting the shaved area, place the animal under a microscope and begin the procedure by making a small longitudinal incision about 1.5 centimeters over the desired spinal level.
Expose the back musculature by carefully retracting the subcutaneous connective tissue. Then carefully separate the paravertebral muscles from the spinal column at the desired level. Make as few incisions as possible to detach the muscles from both sides of each spinous process.
Afterward, scrape the detached muscles away from the laminate surface. Use a small vessel cauterizer to control any minor bleeding if necessary. Continue scraping the muscles until you have sufficiently exposed the underlying laminate, and you can clearly identify the one that you are going to remove.
To prepare the sites of entry for the spinal clamps on each side of the spinal column, insert the tips of a pair of curved scissors between the paravertebral muscles and carefully displace the muscular tissue to create four pockets where the clamps will be inserted. Lift the spinal column with the straight tooth tip forceps to allow safe insertion of the curved blunt tip scissors under the lamina without touching the spinal cord. Then perform the laminectomy by slowly cutting the bone on one side and then on the other side, use a cotton swab or insert a precut small gel foam absorbable gelatin sponge piece next to the incision to limit minor bleeding.
After that, use the preheated sterile artificial cerebral spinal fluid to wash the tissue. A spinal stabilization device was custom designed with the main components of the Nagate STSA complex spinal cord clamps, and the Nagate MA six N head holding adapter. A stainless steel base plate was made to hold two naga parts in alignment so that the animal's head is supported while its spinal column and tail are clamped.
Keep in mind that the entire device should fit under the microscope lens on a lowered microscope stage to stabilize the animal. First, place it on an elevation pad on the base plate of the spinal stabilization device. After securing the head in the head holding adapter, place the spinal clamp of the SDSA device along the anterior posterior axis of the animal in the pre-prepared pockets, flanking the laminectomy.
Place the two clamps at an angle of around 45 degrees relative to the animal's ROS cordal axis to allow enough space for lowering the water immersion lens over the exposed spinal cord. Then place the third clamp at the base of the tail so that the animal's body can be suspended after removal of the elevation pad in the imaging experiment. Next, build a small well of gel seal around the exposed spinal cord to facilitate the maintenance of the spinal cord in A CSF.
Then remove the elevation pad when you are ready to begin In vivo imaging, bring the stabilization device to the preheated microscope chamber and place it on the lowered microscope stage. Make sure to align the laminectomy with the lens. Lower the water immersion lens carefully into the A CSF solution, and make sure that it does not touch the spinal clamps or the gel seal.
Use epi fluorescence to identify the area of interest and focus on it afterwards. Switch to laser scanning mode and perform in vivo imaging using the appropriate two photon laser excitation, wavelength, dichroics, and band pass filters for the fluor's present in the image tissue. At the end of the imaging experiments, remove the animal from the spinal stabilization device.
Carefully wipe the gel seal from the area around the laminectomy and clean it. Well suture the back muscles over the laminectomy. Then suture the skin incision and swab it with Betadine shown here as a projection of Zack images as they were required in vivo and subsequently projected along the Z axis without image alignment.
The image is showing highly dense GFP positive microglial cells in green, enclosed proximity with blood vessels in red in the spinal cord. Here is a high magnification image of a single microglial cell attached to the wall of a blood vessel showing in detail some of its processes extended around the vessel and some towards the spinal cord parenchyma shown Here is an example of repetitive in vivo imaging of the same YFP labeled axonal segments as they were relocated and reim imaged in the spinal cord of the same YFPH line mouse on two different sessions five days apart. This is another example of repetitive imaging showing the same vascular structures and microglial cells imaged in vivo in the spinal cord on two consecutive days.
Here is a representative time-lapse sequence that was acquired in vivo from the spinal cord. This sequence shows in detail the dynamic behavior of the finer processes of microglial cells in green and their interactions with the vasculature in red. Over time, the raw images were only corrected for background noise, brightness and contrast, and the time-lapse movie was constructed by ze rejecting sequentially image stacks without image alignment.
Averaging Aussie selection of individual planes, blood vessels go through a similar range of Z planes as the images of microglia. The depth of the image volume is 38 microns. This is a time-lapse sequence showing the stability of the imaging method at the level of a single Z plane.
Fast acquisition of the same single Z plane in the spinal cord of a CX three, CR one GFP heterozygous mouse anesthetized with the KXA mix and placed on the spinal stabilization device demonstrates minimal image displacement between consecutive frames. Notably, the rate of this sequence was one frame per second, which is faster than the breathing rate of the mouse. The minor residual displacement is probably due to the animal's heartbeat.
While attempting this procedure, it is important to remember to limit the number of incisions to the minimum and try to limit any minor bleeding as soon as it occurs. Now after watching this video, you should have a good understanding of how to perform stable in vivo imaging in the mouse power cord using two photo microscopy.
