Neuroscience
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Quantification of Cerebral Vascular Architecture using Two-photon Microscopy in a Mouse Model of HIV-induced Neuroinflammation
Chapters
Summary January 12th, 2016
This paper describes a method by which the vascular architecture in the brain can be quantified using in vivo and ex vivo two-photon microscopy.
Transcript
The overall goal of this procedure is to quantify changes in the cortical capillary networks after prolonged exposure to the HIV virotoxin, tat. This method can help to examine key changes in cortical capillary morphology which may contribute to the pathogenesis of HIV associated neurocognitive disorders. The main advantage of this technique is that it allows for physiologically accurate quantification of multiple capillary morphological parameters.
Though this method can provide insight into the pathogenesis of HIV associated neurocognitive disorders. It can also be applied to other diseases in which vascular changes occur such as Alzheimer's disease. Begin this procedure by applying artificial tear gel to the eyes of an anesthetized mouse.
Next, shave its head using an electric razor. Then, apply povidone iodine solution to sterilize the scalp, and allow it to dry. Under a light microscope, remove the scalp of the animal, to completely expose the parietal bones, the caudal frontal bones, and the bregma point.
Apply a small quantity of a ten percent ferric chloride solution to the skull, in order to dry the membrane for easy removal. Subsequently, remove the dried membrane, by gently scraping the skull with the forceps. Then, apply a thin layer of glue around the head plate window.
Gently press the head plate against the skull of the mouse, to keep the area of interest in the center of the window. Afterward, apply a drop of dental cement to the head plate, to polymerize the glue. Next, apply a thin layer of glue along the edge of the head plate window to create a reservoir to hold the saline.
Once the glue has dried, screw the head plate into the mouse head plate harness. Remove any glue from the head plate window using a drill bit attached to a micro-torp drill, set at 6, 000 rpm. Stop every 10 to 15 seconds to prevent overheating of the mouse cranium.
After that, using a new drill bit, begin to thin the skull using a micro-torp drill set at 4, 000 rpm. Move the drill gently across the skull without direct downward pressure. Once the skull is completely thinned, detach the mouse from the holder and place it on its back.
Gently tape down both hind limbs to clearly expose the medial thighs. Then remove the hair over both medial thighs and disinfect the surgical site by covering the thighs with povidone iodine. Next, gently remove the skin on the medial right thigh, above the femoral vein and artery.
Apply approximately three to five drops of 0.9%saline to the surgical site. Then, separate the femoral vein from the artery, by bluntly dissecting into femoral neurovascular bundle. Now, place two three centimeter pieces of surgical suture, underneath the femoral artery, approximately one centimeter apart.
Twist the upper suture clockwise to create a vascular tourniquet that will help to prevent excessive blood loss during catheterization. Subsequently, make a small incision in the femoral artery using spring scissors, where the catheter will later be inserted to monitor the mouse's physiological parameters. In this procedure, remove the skin on the medial left thigh of the mouse.
Next, locate the femoral vein. Using a one milliliter syringe and a 30 gauge needle, draw up 130 microliters of the fluorescent dye solution. Inject 100 microliters of the dye slowly into the femoral vein.
After removing the needle, apply steady gentle pressure to the injection site to stop any bleeding and allow the dye to circulate for five minutes. Then, close the surgical site with sutures. Carefully flip the mouse onto its stomach and place it into the mouse head plate harness.
For in vivo two photon imaging, move the surgical apparatus to the two photon microscope and make sure to maintain the animal's anesthesia level. Next, place a small amount of 0.9%saline into the head plate reservoir and lower microscope objective so that it comes into contact with the saline. Then, locate the area of interest using the bright field viewing objective.
Afterward, begin two photon imaging. Locate a capillary bed on the view screen and magnify this area using the optical zoom, too. Acquire images of the capillaries using the two photon imaging software.
For ex vivo two photon imaging, make an incision in the scalp from the interparietal bones to the frontal bones of the decapitated mouse. Secure the skin to the sides of the skull with the index finger and the thumb, place the extra fine scissors underneath the medial interparietal bone and cut the skull along the sagittal suture until approximately three millimeters after the bregma point. Then, separate the skull from the brain and carefully remove any meningees from the surface of the brain with the forceps.
Gently slide the forceps under the brain to free it from the skull. Afterward, place the brain in a brain matrix, specific for mice, and wash it with drops of ACSF. Next, remove a millimeter thick coronal brain section.
Place the brain section onto a concave glass slide containing ACSF, with the most anterior part of the section facing upwards. Then gently cover the brain slice with a glass cover slip. Transfer the slide to the microscope stage and place a small quantity of 0.9%saline on the cover slip.
Lower the microscope objective until it comes in contact with the saline. Subsequently, locate the mid-line of the brain using the bright field objective. Now, begin two photon imaging, and locate the mid-line again using the 25x objective.
Accomplish this by looking for the longitudinal fissure at the cortical surface of the coronal section. Then, place the right edge of the imaging screen on the mid-line and move the viewer screen laterally over three complete frames. Locate the depth at which capillaries are barely visible on the view screen.
Then, lower the plane of focus for an additional 20 micrometers to determine the top of the z-stack. Set the image thickness to one micrometer. Lower the view screen for 100 micrometers and adjust the laser power throughout such that less than one percent of the pixels are over-saturated.
Finally, acquire the z-stack. In this figure, after the brain slice has been prepared, an imaging area at approximately 1.5 millimeters from the mid-line is located. A 100 micrometer z-stack produces a three dimensional image used for analysis in the Amira Analytical Software.
The capillary network is then manually traced by placing nodes at the beginning and end of a capillary or any location where one capillary branches into another. This produces a fully skeletonized image from which the morphological parameters are automatically extracted. The number of capillary nodes, segments, mean segment length, and the total segment length can be extracted using two photon ex vivo imaging of mouse brain slices.
Here, a two dimensional image of the capillary networks is produced by in vivo imaging, in which the capillary diameter can be extracted and the total capillary volume can be calculated using the capillary diameter and the total segment length obtained from the ex vivo imaging. Once mastered, this procedure can be completed in three and a half hours if it is performed properly. While performing this procedure, it's important to move quickly while maintaining a steady level of anesthesia as isoflurane can cause vasodilation of cortical capillaries, thus skewing the data.
During this procedure other capillary parameters may be obtained, such as red blood cell velocity or red blood cell flux, which can provide a deeper understanding of the pathogenic changes seen in HIV associated neurocognitive disorders and other neurodegenerative diseases. Good luck with your experiments.
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