Prep

1. Check equipment, tools and reagents:

   • Equipment: drill bit (#56 for rats), nose cone (rat or mouse), ear bars (rat or mouse), picospritzer

   • Tools (place these on a new piece of sterile cloth beside the stereotax): sterile cotton tips, tweezers, scalpel, hemostats (x2), cauterizer, scissors, non-absorbable sutures

   • Reagents: isoflurane, eye lubricant, hair remover (e.g., Nair), alcohol wipes, iodine (betadyne), hydrogen peroxide, ketoprofen (rat or mouse)

   

   • Virus Tools: virus, dye (optional), pipets (1-5 uL disposable micro-pipets, Fisher Scientific Cat. No. 21-164-2A), pipet holder, pipet syringe
     
     

   

2. Make pipets

   • Center a pipet in the pipet puller and choose program #4, then press start. For DV coordinates greater than 5 mm, use the vertical puller in the Guyenet lab instead.
     
   • Mark each pipet with a sharpie, placing a mark at every 6 mm (corresponds to about 300 nL).
     
   • Make one pipet for each animal to be injected and make a few extras just in case.
     
     

3. Prepare the coordinates to be used for injections. Use the Angled Injection Calculator for angled injections.

4. Prepare virus

   • Take out aliquots from the -80 degrees freezer and place them on ice.
     
   • Update the virus spreadsheet (use the beenhakkerlab Google account to access this).
     

   • Dilute virus with dye if desired. Adjust the volume to inject accordingly if virus is diluted.

   • Note: Only thaw and load pipet with virus right before the injection!

5. Check charcoal filter

   • Weigh the charcoal filter on the red scale. Record new mass with name and date. Don't use the scales in the chemicals area because they will max out.
     
   • Replace with a new filter (and label the old filter with colored tape) if it's more than 50 g higher than initial mass.
     
     

6. Check oxygen and nitrogen tank levels. Replace if necessary.

7. Check isoflurane levels and refill using a nozzle if necessary.

8. Cover the heating pad with a new sterile cloth.

9. Turn on the master switch of the power cord below the stone table. This should turn the stereotax, the microscope and the heating pad on.

10. Turn on oxygen tank and set air flow to 2-2.5 for adult rats or ~1 for rat pups. Turn on the butterfly lamp.

Anesthesia

1. Initial knockout: Open the air flow to box, place animal in anesthesia box, and set isoflurane level to 5.

2. Open air flow to cone. Flush out lines with oxygen and close the air flow to box. Don't close the air flow to box before opening air flow to cone!

3. Transfer animal to nose cone, and reduce isoflurane level to 2-3. Monitor the animal's breathing pattern and adjust the isoflurane level accordingly throughout the surgery.

4. Apply eye lubricant to both eyes with a cotton tip.

5. Use tweezers to help position the animal's jaws on the bite bar. For smaller animals, tape over extra space in the nose cone to prevent isoflurane from leaking out.

6. Pinch toe to check consciousness, then insert ear bars. Use tweezers to help position the ears so that the bars go in the ear canal. Tug on the tail slightly to see if the ear bars are tight enough. When the ear bars are placed correctly, the skull should not move upon downward pressure.

Surgery

1. Remove hair with Nair (may take a few applications). The area of interest should be posterior to the eyes. In pups the skull sutures should already be visible at this stage.

2. Sterilize the scalp three times alternating with iodine (betadyne) and alcohol wipes.

3. Before cutting, check the consciousness level and adjust the isoflurane level if necessary.

4. Using a scalpel, firmly cut down the midline of the scalp.

5. Peel back scalp and periosteum and use hemostats to hold them out of the way.

6. In adults, cauterize bleeds. In pups, just apply pressure with a Kimwipes.

7. Clean skull with hydrogen peroxide. The sutures should become even more apparent.

8. Clean the area with cotton tips.

9. Attach the drill to the stereotax. Angle the arm if angled injections are to be performed.

10. Move the drill bit head to bregma and zero all coordinates on the stereotax.

11. Drill all holes for where the virus is to be injected.

12. Thaw a virus aliquot. Break the pipet tip slightly by poking through a Kimwipe. Using a pipet syringe, pull out enough virus for this animal. Usually one injects around 300 uL per site.

    Note: Make sure to equilibrate to atmospheric pressure before taking the filled pipet out of the tube.

13. Attach the filled pipet to the picrospritzer, then attach it to the stereotax arm with the holder.

14. Turn on the nitrogen tank.

15. Position the pipet with the stereotax:

    • Move the pipet tip to bregma and zero all coordinates on the stereotax.
      
    • Raise the pipet and move it to the correct ML and AP coordinates.
      
    • Lower the pipet just enough to see a reflection on the dura, then zero the DV coordinate.
      
    • Lower the pipet while wiggling up and down to the site of injection. This allows the brain matter to fully surround the pipet because it gets pushed out of the way upon insertion.
      
      

16. Inject with the picospritzer:

    • Turn the regulator knob on the Eject panel all the way down on the picospritzer.
      
    • Press the start button while looking at the fluid level in the pipet. Gradually increase the pressure until the fluid level drops about 1-2 mm at a time.
      
    • Inject once every 30 seconds. One can use the Echo behind the stereotax to set timers.
      
    • Make sure to leave the pipet in place at the injection site for at least 30 seconds before taking it out to prevent backflow.
      
      

17. After all injections, remove the pipet and turn off the nitrogen tank.

18. Use tweezers to close the scalp. With the aid of hemostats, tweezers and scissors, suture using whatever method you like (e.g., instrument tie).

19. Inject 0.1-0.15 mL ketoprofen in the back subcutaneously and massage.

Post-Op

1. Turn off the oxygen tank. Turn off the butterfly lamp.

2. Clean tools with alcohol and DI water.

3. Turn off the master switch to the power cord.

4. Place animal on its side in a clean cage.

5. Fill out a surgery card and wait for it to wake up before taking it to the vivarium. Add a water bottle if necessary and place the cage in the biohazard room B033.

6. Check on rat everyday for three days following surgery updating the surgery card each time.

7. Remove cage from biohazard room after three days and return it to our normal rat room.

8. After tools are dried, either autoclave them or place in the Germinator.