Generating a Double Humanized BLT (Bone-Marrow, Liver, Thymus) Mouse Model

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Mice housing and maintenance

1. Purchase 6-8-week-old NSG mice (NOD.Cg-Prkdc^scidIl2rg^tm1Wjl/SzJ).
2. House the mice under SPF conditions with air exchange, prefilters, and high-efficiency particulate air (HEPA) filters (0.22 μm) in a room with controlled temperature, humidity, and pressure.
   1. House and maintain the mice in autoclaved individual microisolator cages in a rack system capable of managing air exchange with prefilters and HEPA filters (0.22 μm).
3. Perform all procedures and manipulations of the mice in a Class II Type A2 biological safety fume hood that has been pre-treated with 70% ethanol. Before working in the animal room, shower and change into clean scrubs, Tyvek suit, boot covers, hair cap, face mask, and gloves. Wear a surgery gown over the Tyvek suit during surgical procedures.         
   NOTE: Ultraviolet (UV) light decontamination is also preferred prior to procedures.
   1. Autoclave all instruments and reagents if possible, and then disinfect them prior to transferring them to the fume hood.
   2. During procedures, remove the animals' individually ventilated cages from the rack and disinfect them prior to transferring them to the fume hood.
4. Feed the mice an irradiated diet and provide autoclaved water. Provide irradiated food and supplement with additional irradiated food as needed. Change autoclaved water weekly or as needed.   
   NOTE: Autoclaved acidified water can also be used. The irradiated diet provided had a shelf-life of 6 months after manufacture.

2. Generation of humanized bone-marrow, liver, and thymus (BLT) mice

NOTE: The generation of hu-BLT mice has been described previously.

1. On the day of surgery, give the mice whole-body irradiation at a dose of 12 cGy/g of body weight.
2. Prepare the mice for surgery.
   1. Give each irradiated mouse a mixture of Ketamine at the working concentration of 100 mg/kg and Xylazine at the concentration of 12 mg/kg, which ranged from 130-170 μL per mouse based on body weight by intraperitoneal (IP) injection for anesthesia. To prepare 3 ml of the Ketamine and Xylazine mixture, add 0.27 mL of Ketamine at the stock concentration of 100 mg/mL and 0.03 mL of Xylazine at the concentration of 100 mg/mL to 2.7 mL of sterile saline and mix well. Disinfect the skin with 70% isopropanol prior to injection.
   2. Give each irradiated mouse Buprenorphine 1 mg/kg of body weight (half-live 72 h, SR-LAB) by subcutaneous injection for long lasting pain management.
   3. Give each irradiated mouse 100 μL (858 μg) of Cefazolin by IP injection for preoperative antibiotic prophylaxis.
   4. Shave the hair of the mice using electric hair clippers around the left lateral and medial side of the mouse at the later surgical site used to expose the left kidney.
   5. Verify proper level of anesthesia by pedal reflex (firm toe pinch).
   6. Give isoflurane gas at 3-5% if additional anesthesia is needed at any point during surgery.
   7. Apply ophthalmic ointment to both eyes to prevent corneal desiccation. Apply ear tag if needed.
3. Perform surgery to implant the liver and thymus tissues into the left kidney capsule.
   1. Disinfect the skin at the surgical site by applying iodine scrub starting from the center of the surgical site and moving towards the outside in a circular manner. Repeat this process with 70% isopropanol and a third time with iodine.
   2. Using forceps, first load one human fetal liver tissue fragment into a trocar. Then using forceps, load one human fetal thymus tissue fragment into the trocar. Then using forceps, load another human fetal liver tissue fragment into the trocar. Tissues should be cut to 1-1.6 mm³ to fit the inner dimensions of the trocar.
   3. Use forceps to lift the skin and use scissors to make a small cut longitudinally. Extend the cut to 1.5-2 cm in the left side of the mouse.
   4. Use forceps to lift the muscle layer and use scissors to make a small cut longitudinally. Extend the cut as needed to expose the kidney.
   5. Expose the kidney by gently grasping the fatty tissue surrounding the kidney. Do not directly touch the kidney parenchyma.
   6. Make a 1-2 mm incision at the posterior end of the kidney capsule using a scalpel.
   7. Slowly insert the preloaded trocar through the incision parallel to the long axis of the kidney and release the tissues between the kidney capsule and kidney.
   8. Carefully return the kidney and bowel to their normal position. Use absorbable 5/0 P-3 (P-13) sutures with 13 mm 3/8 circle needle to close the muscle layer and surgical staples to close the skin.
4. After surgery, put the mouse into a clean autoclaved microisolator cage for recovery.
   1. To minimize heat loss during post-surgical recovery, put the cage containing the mice on a heating pad that is connected to a water pump that warms and circulates the water.
   2. Monitor the mice until they have regained sufficient consciousness to maintain sternal recumbency.
5. Within 6 h of surgery completion, via the tail-vein, inject CD34+ hematopoietic stem cells isolated from human fetal liver tissue.
   1. Warm up the mice with a heat-lamp. Disinfect the tail with 70% isopropanol and then inject 1.5 to 5 × 10⁵ stem cells/200 μL into the tail vein.
   2. Stop any bleeding from the injection and return mice to the microisolator cage and the microisolator cage to the cage rack.       
      NOTE: Post-surgery mice are typically housed together, five per microisolator cage, during and after recovery. However, mice are only housed together if they all received surgery on the same day.
6. Check the mice daily. Carefully monitor the surgical staples and replace them as needed. Closely monitor the mice for any sign of infection or discomfort. Supply autoclaved food on the floor of the microisolator cage for a few days post-surgery.
7. Remove the surgical staples 7-10 days after surgery. Give isoflurane gas at 3-5% to anesthetize the mice. Carefully remove the staples and then apply antibiotic and pain-relieving ointment onto the site.
8. Allow 9-12 weeks for reconstitution of human immune cells, then collect peripheral blood from the medial saphenous vein from each of the humanized mice.
   1. Restrain conscious mice using an appropriately sized plastic cone restraint with an opening near the head of the mouse for breathing and an opening near the back of the mouse to isolate one leg. Put the mice into the restraint cone head first and then gently pull one leg through the leg opening.
   2. Spray the medial side of the isolated leg with 70% isopropanol, then spread antibiotic and pain-relieving ointment onto the same site. 
      NOTE: The ointment helps to reveal the location of the vein without the need for hair removal and also assists in blood droplet formation.
   3. Using a 25-gauge needle at a 90° angle, puncture the vein and collect 50-100 μL of blood using an ethylenediaminetetraacetic acid (EDTA) coated blood collection tube. Stop the bleeding by applying pressure to the site with sterile gauze. Once the bleeding has stopped, return the mice to their cage.
      NOTE: The maximum blood volumes collected are typically 50 μL per week or 100 μL every two weeks.
   4. Use the collected peripheral blood to test the level of human immune cell reconstitution using flow cytometry with antibodies for hCD45, mCD45, hCD3, hCD4, hCD8, hCD19.

3. Antibiotic treatment

1. Prior to antibiotic treatment, collect pre-treatment fecal samples. Move the mice to a new autoclaved microisolator cage.
2. Prepare a fresh cocktail of broad-spectrum antibiotics daily.
   1. Prepare 250 mL of water containing freshly prepared Metronidazole (1 g/L), Neomycin (1 g/L), Vancomycin (0.5 g/L), and Ampicillin (1 g/L) for each microisolator cage of mice.  
      NOTE: Use autoclaved or sterile water for the antibiotic supplemented drinking water.
   2. Add 9.2 g of grape sugar sweetened drink mix to 250 mL of antibiotic supplemented water. 
      NOTE: The use of grape sugar sweetened drink mix masks the bitter taste of the antibiotics and helps prevent dehydration in the mice.
   3. Change the antibiotic and grape sugar sweetened drink mix supplemented water and place the mice in a new autoclaved cage daily. 
      NOTE: Mice are coprophagic and changing the cages daily prevents the mice from re-inoculating themselves with gut bacteria.
   4. For maximal engraftment of the human-like gut microbiome, provide antibiotics for 14 days. During antibiotic treatment, monitor the mice for weight loss and dehydration. Weight loss is expected during the first 3-4 days and plateaus after that for the duration of the treatment. If dehydration occurs, give the mice saline or Ringers Solution via intraperitoneal injection.
      NOTE: Other gut microbiome humanization protocols call for oral gavage of antibiotics. While effective at depleting the murine gut bacteria, we found that the less invasive method of adding antibiotics to the drinking water put less stress on our humanized mice and led to better outcomes.

4. Donor samples and fecal transplant

1. Prepare human donor fecal samples.
   1. Use properly prepared sources of fecal microbiota transplant (FMT) material for fecal transplant into the humanized mice.
   2. Thaw FMT preparations and aliquot them under anaerobic conditions in an anaerobic chamber.
   3. If desired, at this step mix equal parts of the fecal samples together to create an unbiased "human" sample.
   4. Keep freezing and thawing of the FMT material to a minimum, if aliquoting or mixing of samples is not needed, only thaw immediately before the procedure.
2. Human fecal transplant
   1. After completion of 14 days of antibiotic treatment, change the drinking water to autoclaved water and move the mice into a new autoclaved cage. Stop daily cage changes and implement a once every 1-2 weeks cage changing schedule.
   2. Give two fecal transplants at 24 and 48 hours post-cessation of antibiotics.
   3. 24 h after antibiotics treatment, thaw the needed amount of FMT material and give each mouse 200 μL of FMT material via oral gavage. Repeat the procedure again at 48 h post antibiotics.
   4. Spread any remaining or leftover thawed FMT material on the fur of the humanized mice or onto the cage bedding.