Imaging Glioma Initiation In Vivo Through a Polished and Reinforced Thin-skull Cranial Window

The overall goal of this procedure is to visualize initial stages of glioma initiation from single injected glioblastoma cells in vivo. This is accomplished by first thinning, an area of mouse skull down to a thickness suitable for in vivo imaging purpose. Next, the thin skull area is polished.

Then GFP labeled glioblastoma cells are injected into the mouse brain and the cranial window is sealed for in vivo imaging. Finally, the injected glioblastoma cells are imaged daily to visualize glioma initiation from injected glioblastoma cells in vivo. Ultimately, results can be obtained that show in vivo glioma initiation from a single injection of glioblastoma cells and its relationship with vascular endothelial cells through dual color in vivo two photon microscopy.

This method can help answer key questions in cancer stem cell field, such as the critical role of interaction between glioma stem cells and vascular endothelial cells and glioma initiation in vivo, demonstrating the procedure will be under lapier manager of JAK Surgical Services and postal in my laboratory To prepare for surgery in an autoclave, steam sterilize all surgical tools, including the dental drill bit after the mouse has been anesthetized with ketamine and xylazine and administered analgesia. Check the level of anesthesia by performing a toe pinch using a small electric shaver. Remove the fur from the dorsal head in a roughly triangular area, about three millimeters, coddle to the nares and extending coddly to the cervical vertebrae.

Apply ophthalmic ointment to the eyes. Place the mouse in ventral recumbent and use surgical iodine followed by 70%ethanol. To disinfect the skin, place the animal on a cotton pad and transfer it to a heating pad.

To maintain a body temperature of about 37 degrees Celsius, apply a topical analgesic to the periosteum and the exposed tissues of the wound. Using a scalpel blade, remove the periosteum by gently scraping the skull. This will help the cyanoacrylate glue adhere to the bone.

Next, use a marker to outline the cortical area of interest, making sure to avoid suture lines to avoid damaging the underlying large vessels. Then apply a thin layer of cyanoacrylate adhesive to the wound margins to prevent the seepage of serosanguinous fluid and to the skull except in the area of interest before sealing the skull with light activated dental cement, treat all exposed skull except the area intended for thinning with primers and then bonding agent. Once the bonding agent has dried, cover the skull with dental cement to stabilize the mouse's head.

For surgery and imaging, embed a number double zero dash 90 hex nut into the dental cement at a distance from the area to be thinned. Once the dental cement preparation is completed, use visible light to cure it. Then using a number zero zero dash 90 screw mount, a headpiece holder from the stereotactic device to the hex nut.

Next, place the mouse's head under a stereo microscope and apply a drop of room temperature, 0.9%sterile saline to the skull area to be thinned. Use a high speed micro drill to thin a circular area of the skull over a region of interest. To avoid thermal injury to the underlying cerebral tissue, intermittently pause drilling and apply saline.

The mouse skull comprises two thin layers of compact bone, sandwiching of thick layer of spongy bone. Use the drill to remove the top thin layer and most of the spongy bone. After the majority of the spongy bone has been removed, the remaining cavities within the spongy bone can be seen under the dissecting microscope indicating that drilling is approaching the internal compact bone layer.

At this stage, skull thickness should still be more than 50 microns. Carefully continue skull thinning to obtain a very thin and smooth preparation. To further thin the skull, use a custom made silicone whip and diamond paste to polish it for about 10 minutes.

Continue to polish the skull with fine-grained tin oxide for an additional 10 minutes. Use saline to flush away the tin oxide until the thin skull appears clean. To perform an injection, use a 26 gauge syringe needle to create a small opening on one side of the polished window under the microscope.

Use the micro manipulator to lower the tip of the pipette through the hole in the cranial window and into the brain. 100 to 200 microns deep. Then inject one microliter of cell suspension at a rate of 0.1 microliters per minute.

After removing the needle to seal the cranial window, dry the skull and apply a small drop of clear cyanoacrylate glue and a piece of three millimeter. Number one, cover slip. Use cyanoacrylate glue to seal the space between the cover slip and adjacent dental cement.

After surgery, inject the mouse subcutaneously with one milliliter of warm sterile saline and provide supplemental heat to maintain body temperature until fully recovered from anesthesia. Administer subcutaneous injections of analgesic daily for five days after surgery, and observe the mouse daily to ensure that the wound is healing properly and that the mouse is eating, drinking, and behaving normally to image. After anesthetizing the mouse with ketamine and xylazine immobilize it using a custom stereotactic frame under the microscope.

A successful ports cranial window surgery allows the cranial window to remain clear for weeks to months. This figure shows the vasculature between a port's cranial window as visualized using back scattered light. These vasculature images were used as landmarks for locating the same brain areas for repetitive imaging.

To visualize both the vascular endothelial cells and GBM cells, mice were generated with vascular endothelial cells labeled with the red fluorescent protein TD tomato. After the port's craniotomy procedure, GFP labeled GBM cells were injected into their brains, and in vivo two photon imaging was performed.Shown. Here is an in vivo example of glioma initiation from injected GBM cells near the vasculature.

This figure shows GFAP staining of cortical sections from a mouse without port surgery, a mouse seven days following port surgery, and a mouse seven days following port surgery with saline injection. It is evident from the images that the mouse without surgery and the mouse with port surgery do not have gliosis. Whereas following saline injection, gliosis is observed in the brain tissue of the mouse due to pipette penetration and saline injection in the brain.

This result demonstrates that gliosis does not occur under the port's cranial window. Providing independent validation that the port's cranial window described here does not lead to the gliosis that is typically associated with an open skull chronic cranial window. Here, high resolution imaging of dendritic spines are shown from the same mouse on the day of surgery, day zero and 32 days after the port's window creation.

Demonstrating that the port's window is also an excellent choice for high resolution in vivo imaging. After watching this video, it should have a good understanding of how to image glioma initiation by injecting glioblastoma cells through a polish and reinforce the cranial window.