Constructing a Humanized Mouse Model with Engineered Immunity against HIV

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines. All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Construction of Humanized Mice Engineered with CD4 Chimeric Antigen Receptor

1. Processing Fetal Thymus and Isolating CD34+ HSCs from Fetal Liver
   1. Thymus Processing
      1. Gently wash the thymus in Phosphate-buffered saline (PBS), pH 7.4, in a 15 ml conical tube. Repeat the wash step 3 – 4 times.
      2. Add 7 ml Roswell Park Memorial Institute (RPMI) media + 10% fetal bovine serum (FBS) + penicillin/streptomycin (Pen/strep). Decant everything into a sterile 100 mm Petri dish.
      3. Use scalpels to cut the thymus into small pieces of about 1 mm². Place every single thymus piece in a single well on a 96-well plate. Use curved blunt forceps when transferring thymus pieces to the 96-well plate.
         1. Add a small amount of media (100 – 200 μl) to all the wells so the tissue does not dry. Visualize under a microscope (thymi have lobes and should look like sacks of cells).
            NOTE: Discard any pieces that look questionable in any way; there is often connective tissue, and this should not be implanted.
      4. Remove the confirmed thymus pieces and place all of them into a T25 cell culture flask. Add 7 ml RPMI media supplemented with 10% FBS and 450 μg/ml of piperacillin/tazobactam and amphotericin B. Gently rock flask to mix. Culture the flask overnight at 37 ºC/5% CO₂.
         NOTE: This step is required to prevent bacterial contamination of the tissue.
      5. (Optional step) Freeze the thymus for future use. Equilibrate the chunks in 90% Human AB Serum with 10% Dimethyl sulfoxide (DMSO) for 10 min. Freeze them at 1 ºC/min to -50 ºC, then rapid cooling to -150 ºC. When ready to thaw, rapidly thaw in a 37 ºC water bath and gently wash 3x in RPMI complete media without DMSO.
   2. Liver Processing
      1. Gently wash the liver in PBS in a 50 ml conical tube. Repeat the wash step 3 – 4 times.
      2. Add 10 ml Iscove's Modified Dulbecco's Media (IMDM) to the 50 ml conical tube. Decant everything into a 100 mm sterile Petri dish.
      3. Homogenize the liver tissue using two scalpels. Cut the liver into small pieces of about 3 mm². Cut out and discard any white connective tissue.
      4. Use a 10 ml syringe fitted with a 16-gauge blunt needle to take up the liver pieces and media. Then, transfer to a 50 ml conical tube.
      5. Resuspend the media and tissue suspension and expel 5-7 more times to homogenize the tissue completely.
      6. Prepare 10 ml IMDM media supplemented with enzymes: 500 U/ml collagenase, 2,400 U/ml hyaluronidase, and 300 U/ml DNase as well as 450 μg/ml piperacillin/tazobactam and amphotericin B. Filter the media via a 0.22 µm filter then add the media to the liver suspension.
      7. Cap the 50 ml conical tube containing the liver suspension and seal tightly with a self-sealing film such as Parafilm to prevent leaks. Rotate in a tube rotator in the incubator at 37 ºC for 90 min.
      8. Filter the digested cell suspension through a 100 µm cell strainer into a fresh 50 ml tube.
         NOTE: Add PBS to the suspension to increase the total volume to 50 ml. Split this into two 50 ml tubes, each containing 25 ml of cell suspension.
      9. Slowly and gently underlay the cells in each tube with 10 ml density centrifugation media (e.g., Ficoll). Spin at 1,200 x g for 20 min without brake. Note: all centrifugation mentioned in this protocol is done at room temperature (25 ºC).
      10. Carefully remove the interface (i.e., buffy coat) from each tube and transfer to two separate 50 ml tubes. Bring the volume of each tube of interface up to 50 ml with PBS. Spin at 300 x g for 7 – 10 min. Aspirate supernatant carefully.
      11. Combine the two pellets. Wash three more times with 50 ml PBS containing 2% FBS. Spin at 300 x g for 7 – 10 min each while carefully aspirating the supernatant.
      12. Resuspend the pellet in 50 ml RPMI media + 10% FBS. Count cells using a hemocytometer before proceeding to cell sorting.
      13. According to the manufacturer's protocol, sort CD34+ cells immediately using a CD34 sorting Kit (e.g., CD34 micro-beads).
          NOTE: Alternatively, cells can be cultured overnight in RPMI media + 10% FBS at 1 million/ml.
      14. Save both CD34+ and CD34- fractions.
          NOTE: At this step, CD34+ and CD34- cells can be frozen using Bambanker freezing media or other freezing media. Freeze 1 ml of 4 – 6 x 10⁶ CD34+ cells per tube and freeze 1 ml of 40 – 60 x 10⁶ CD34- cells per tube.
2. Transduction of CD34+ Cells
   1. Calculate the number of transduction wells required for a 6-well-tissue-culture plate. One well can be used to transduce up to 8 x 10⁶ cells. For each Bone-marrow/Liver/Thymus (BLT) mouse, 0.5 x 10⁶ CD34+ cells will be implanted along with CD34- cells and thymus underneath the kidney capsule, and 0.5 x 10⁶ CD34+ cells will be injected intravenously. The number of CD34+ cells to use is determined by the number of mice (1 million cells per mouse).
   2. Coat the needed number of non-tissue culture-treated 6-well plate wells with 1.25 ml of recombinant human fibronectin solution (e.g., Retronectin) (20 µg/ml in PBS) into each well. Cover the plate and allow it to stand for 2 hr at room temperature in a clean biosafety cabinet.
   3. Aspirate fibronectin solution from wells and add 1.25 ml of fluorescence-activated cell sorting (FACS) buffer (PBS with 4% FBS) to each well for blocking. Allow the plate to stand at room temperature (25 ºC) for 30 min.
   4. Aspirate FACS Buffer and wash wells once with PBS.
   5. Keep PBS in the coated wells until the plate is ready for use. Store the plate at 4 ºC overnight if not used immediately.
   6. Plate CD34+ cells in Infection Medium (2% Human Serum Albumin in Yssel's Serum-Free T-Cell Medium) in fibronectin solution-coated wells (~2 x 10⁶ cells/ml) and incubate at 37 ºC for 1 hr.
   7. Use a pipette to add lentiviral vectors to the wells at a multiplicity of infection (MOI) between 2 – 10. Gently mix and incubate overnight at 37 ºC.
      NOTE: The titer of the lentivirus vector used should be determined beforehand.
   8. Harvest the cells the following day by gently scraping the bottoms of the wells with a cell scraper. Collect cells and count with a hemocytometer.
   9. To prepare cells for the implant, combine 0.5 x 10⁶ transduced CD34+ cells with 4.5 x 10⁶ CD34- cells per mouse, aliquot into sterile 1.5 ml screw-cap tubes. Spin the cells down at 300 x g to the pellet and aspirate the supernatant. Spin them again at 300 x g, aspirate any remaining supernatant using a P10 pipette, and aspirate very carefully. Keep the dry pellets on ice throughout the study. Note: To ensure the cells are viable, use the pelleted cells and perform the surgery within 2-3 hr.
   10. To prepare cells for injection, spin 0.5 x 10⁶ transduced CD34+ cells per mouse to pellet and aspirate supernatant. Resuspend cells in 100 µl RPMI media per mouse and keep on ice.
   11. To check transduction efficiency, aliquot ~1 x 10⁵ non-transduced and transduced CD34+ cells from each condition and culture in 200 µl cytokine medium (RPMI media with 10% FBS, supplemented with 100 ng/ml human IL-3, IL-6, SCF) in 96-well plate for 5 – 7 days at 37 ºC.
       NOTE: Cells used in this step are not used for surgery but to ensure that transduction is successful and the vector is not toxic for stem cell survival and renewal. Transduction efficiency can be checked by looking for gene expression of the vector (e.g., GFP&CD4) and analyzed by flow cytometry.
3. Tissue Transplants to Construct Genetically Engineered Mice​
   1. On the same day before surgery, total body irradiation of the NOD is performed. Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) immuno-compromised mice with a Cesium-137 Irradiator and a dosage of 2.7 Gy (270Rad).
      NOTE: The NSG mice are severely immunocompromised. Therefore, their housing and maintenance must conform to the highest health standards and be handled by highly trained staff.
   2. Pour thymus pieces and medium from the flask into a 60 mm dish. Pour PBS into another 60 mm dish, which will be used to clean the trochar and keep the kidney wet.
   3. Chill positive displacement pipette tips by placing them in open 1.5 ml sterile screw-cap tubes on ice. Keep on ice with the dried cell pellets and gelatinous protein mixture such as Matrigel.
      NOTE: It is important to keep the gelatinous protein mixture and any tubes or tips that will touch it cold until the implant needle is loaded.
   4. Anesthetize the mice: Weigh mice individually and record weights; ear punch the mice to number them. Inject them intraperitoneally with 15 µl of Ketamine (2.6 mg/ml in saline)/Xylazine (100 mg/ml in saline) per gram of body weight). Put the mice back in the cage and wait for them to be fully anesthetized.
      NOTE: Check the anesthesia level of the mouse by squeezing a paw. If the mouse reflexively flinches, administer 25-50% of the original amount of Ketamine/Xylazine to anesthetize the mouse further. Wait until it does not reflexively flinch to perform the surgery.
   5. Using the Oster clipper (shaver), shave the left side of each mouse from hip to shoulder between the center of the back and the stomach. Subcutaneously inject 0.3 ml of diluted Carprofen (6 mg/kg) into the animal's shoulder or inguinal triangle (leg pit). Using a cotton swab, put a small drop of Artificial Tears onto each eye and lay the mouse on its side back in the cage.
      NOTE: Limit surgery prep to one cage (approximately 4 – 5 mice) at a time.
   6. Flush the cannula of the 16-gauge cancer implant needle (trocar) with PBS. Using a pair of blunt curved forceps, place a piece of thymus from the 60 mm dish into the opening of the cannula with the trocar just inside the opening, then pull back on the trocar to aspirate the tissue into the cannula.
   7. Use a positive displacement pipette and a chilled tip to put 5 µl of cold, gelatinous protein mixture into the tube with a dried cell pellet (CD34+ and CD34- mixture for implanted cells) and gently stir to generate cell suspension. Do not pipette up and down. Pipette the gelatinous protein mixture /cell suspension into the opening of the cannula and slowly pull back on the trocar to load the needle.
      NOTE: It is recommended to have a helper to load the pipette while one manipulates the implant needle.
   8.  Swab the shaved area of the mouse with Povidone-iodine and subsequently wipe down the area with an alcohol wipe three times. Determine the darkest spot under the skin. This indicates the location of the spleen. The kidney is approximately 5 mm dorsal to the spleen. Lift the skin with curved forceps and make a 15 mm long incision with surgical scissors in the skin parallel to the spleen. Then, make a similar cut in the peritoneum layer below. In males, the kidney should be easily visible and can be extruded simply by pressing on the abdomen. You can support the kidney with a hemostat or curved blunt forceps. In females, the ovaries tend to block the kidney from easy extraction. Using a hemostat, pick up the ovary and carefully expose the kidney.
   9. Use the needle-tipped forceps to pluck a tiny hole at the posterior end of the kidney capsule.
      NOTE: Do not use these needle-tipped forceps to handle biohazard materials.
   10. Slide the implant needle into this hole and along the kidney until the opening of the cannula is completely covered by the kidney capsule.
   11. Gently extrude the tissue under the kidney capsule and pull the needle back out. The thymus pieces can be sticky, so use curved forceps to make sure the thymus piece does not come out with the needle.
   12. Lift the peritoneum with the forceps and gently use the hemostat to push the kidney back into place. Tie one double-knotted stitch in the peritoneum using 4-0 vicryl absorbable sutures. Use two Autoclips wound clips to close the skin.
   13. Mix the transduced CD34+ cells set aside for injection and uptake 100 µl (0.5×10⁶ cells) into an insulin syringe. Inject these cells into the mouse through retroorbital vein injection or other routes of intravenous injection. Using a cotton swab, put a small drop of Artificial Tears onto each eye and lay the mouse on its side back in a cage.
   14. Once all the mice have been implanted, confirm that the animals are regaining consciousness and ambulating normally before leaving them.
   15. Post-operational care: Subcutaneously inject 0.3 ml of diluted Carprofen (6 mg/kg) and 1.2 ml of sterile saline into each mouse the day after the surgery. 2 and 3 days post-surgery, subcutaneously inject 1.5 ml of sterile saline into each mouse. Monitor the mice and the incisions for 10-14 days following surgery. Remove the Autoclips and weigh the mice after 10-14 days. NOTE: Mice are sluggish after radiation, and saline injection prevents the animals from dehydrating.
   16. After 8 – 10 weeks, check engraftment by bleeding the mice and performing FACS analysis on the peripheral blood, staining for markers such as CD45, CD3, CD4, CD8, and any genes the vector should express.