Controlled Cortical Impact Model of Mouse Brain Injury with Therapeutic Transplantation of Human Induced Pluripotent Stem Cell-Derived Neural Cells

All experiments described in this protocol were reviewed and approved by the Uniformed Services University Animal Care and Use Committee. 

1. Craniectomy and controlled cortical impact

1. Preparation of the controlled cortical impact device and surgical supplies.
   1. Load a 1 mL slip-tip syringe with 0.5 mL of sterile saline for wound irrigation. Attach a 25 G needle to the syringe to control irrigation.
   2. Prepare a dilute solution of CsA in DMSO to final concentration of 1 mg/mL. Load a second 1 mL slip-tip syringe with 0.5 mL of cyclosporine A (CsA) solution for immunosuppression. Attach a 25 G needle or larger to the CsA syringe.
   3. Attach the controlled cortical impact piston to an arm on a stereotaxic frame and set to an angle of 15°. Attach a 3 mm impactor probe to the piston.
   4. Set the velocity of impact to 1.5 m/s and the impact dwell time to 0.1 s to produce a mild cortical injury.
2. Perform a unilateral craniectomy
   1. Place the mouse in an anesthesia induction chamber connected to an isoflurane vaporizer with compressed oxygen source. Induce anesthesia with ~3% isoflurane at ~0.7 L/min oxygen. Check for the depth of anesthesia by the lack of response to the toe-pinch.
   2. Shave the scalp using electric clippers and wipe away any loose fur.
   3. Place the mouse in a stereotaxic frame with attached anesthetic delivery nose cone. 
      1. Place a warming pad set to 37 °C on the stereotaxic frame under the mouse to maintain body temperature under anesthesia. Fix the head in place with ear bars and a bite bar and orient the head such that the skull frontal bone is horizontal. Maintain anesthesia at ~1.5%-2% isoflurane for the duration of the surgery.
   4. To perform preoperative care and aseptic surgical preparation, apply antibiotic ophthalmic ointment to the eyes using a sterile cotton swab. Apply an iodine-based solution to the shaved scalp area. Remove this with 70% ethanol. Cover the animal with a fenestrated surgical drape so that the top of the head is visible but the eyes are covered.
   5. Make a midline incision (1.5-2 cm) on the scalp using a scalpel or scissors.  Use sterile cotton swabs to clean the wound and to clear the fascia left of the midline at bregma.
   6. Use the impactor probe to identify the craniectomy site.
      1. Set the stereotaxic reference point (X = 0, Y = 0) to bregma.  Adjust the probe laterally to 2 mm left of the midline. Outline a 5 mm diameter circle around the probe using a fine-tip surgically safe marker. Raise and rotate the impactor out of position.
   7. Use the high-speed rotary micromotor kit hand tool to make an open hole in the skull using a round-tip 0.6 mm or 0.8 mm burr drill bit at ~70%-80% maximum speed. Apply light pressure to the skull while drilling along the 5 mm circumferential outline to thin this border.
      1. Do not apply excess pressure while drilling. This can cause cortical injury due to vibration, compression, or accidental penetration. Allow the speed of the drill bit to do the work.
      2. Do not drill in any given spot for too long to avoid excess friction heating of the skull. Irrigate the craniectomy occasionally with sterile saline to remove debris and to reduce heating from the rotary tool.
      3. Pay close attention while drilling over the coronal suture line as these points are vulnerable to hemorrhage.
   8. Use a pair of fine tweezers to remove the skull flap when the craniectomy outline is sufficiently thinned. Grasp the flap medially, and gently lift and pull laterally with a radial motion.
      1. Do not to damage the dura mater when lifting the flap; this can cause severe injury and hemorrhaging.
3. Perform a mild controlled cortical impact injury
   1. Clean the impactor probe with a sterile alcohol prep pad. Move the impactor probe back into position over the exposed cortex. Lower the probe until it touches the dura mater surface. Mark this position as Z = 0.
   2. Withdraw the piston and move to Z = -1.0 mm. Discharge the piston to impact the cortex.  
   3. Quickly raise the piston and move the arm out of position. Apply generous amounts of saline to irrigate the cortex after injury. Rinse the surgery site with saline as needed and suture the scalp incision using simple interrupted stiches with 5.0 silk suture.  
4. Perform postoperative care on the mouse
   1. Discontinue anesthesia. Deliver CsA by subcutaneous injection into scruff at 10 mg/kg dose. Place the mouse in a clean and pre-warmed postoperative cage.
   2. Provide acetaminophen analgesic in the drinking water at 1.0 mg/mL.
      NOTE: Provide analgesia according to the appropriate IACUC standard operating procedure and in consideration of experimental outcome variables (e.g., sedation, neuroinflammation).
   3. Provide moistened chow food in a warmed recovery cage to aid in rehydration and recovery.
5. Proceed from section 2 to section 4 above when performing craniectomy-only (sham) controls.

2. Stereotaxic transplantation of cell suspension

1. Begin cell transplantation procedure roughly 24 h after craniectomy.
2. Prepare the cell transplantation equipment and surgical supplies
   1. Fill a 1 mL slip-tip syringe with sterile saline for wound irrigation. Attach a 25 G needle to the syringe to control irrigation.
   2. Fill a 1 mL slip-tip syringe with CsA solution for immunosuppression. Attach a 25 G needle or larger to the CsA syringe. Refill the CsA syringe as needed between surgeries.
   3. Prepare glass needles from 1.0 mm OD borosilicate glass capillary pipettes using standard methods. 
   4. Use fine tweezers to break the needle tips to approximately a 200 µm diameter. Ensure that the cylindrical shaft of the needle is no longer than 2.5 cm.
3. Calibrate the syringe pump for use with a 10 µL syringe.  Enter a flow rate of 0.2 µL/min to deliver a total volume of 2.0 µL.
4. Prepare human induced pluripotent stem cell (iPSC) suspension.
   1. Perform all cell handling in a cell culture BSL-2 hood using standard sterile handling techniques.
   2. Prepare cell cultures in advance according to standard conditions defined for the cell type. Refer to Lischka et al.¹⁷ as an example.
      NOTE: Experiments shown in this demonstration used various neural phenotype cells derived from hiPSCs.
   3. Gently dissociate the cells into a single-cell suspension using a cell detachment solution, or other preferred enzymatic or chemical means.  
   4. Count cells in suspension, then dilute the suspension to 5 x 10⁴ cells/µL in minimal cell culture medium (e.g., DMEM) in a 1.7 mL flip top test tube.  
   5. Observe the following notes for transplantation:
      1. Maintain the cells in suspension at 37 °C for the duration of procedures.  
      2. Load the syringe only immediately prior to performing intraparenchymal injection (step 4.5 below).
         NOTE: Gravity can cause the cell suspension to settle or to cling to the side of the syringe if laid on its side. This leads to irregularities in the number of cells injected.
      3. Allot ~5 x 10⁵ cells (10 µL suspension) per mouse if performing multiple cell transplantation procedures in one day.
5. Perform stereotaxic transplantation surgery
   1. Place the mouse in an anesthesia induction chamber connected to an isoflurane vaporizer with compressed oxygen source. Induce anesthesia with ~3% isoflurane at ~0.7 L/min oxygen.
   2. Place the mouse in a stereotaxic frame with attached anesthetic nose cone.  
      1. Fix the head in place with ear bars and a bite bar and orient the head such that the skull frontal bone is horizontal. Maintain anesthesia at ~1.5%-2% isoflurane for the duration of the surgery.
   3. Perform preoperative care and aseptic preparation
      1. Apply hydrating ophthalmic ointment containing antibiotic to eyes using a cotton swab.  
      2. Lavage the incision site with sterile saline to clean the site and to loosen sutures. Gently apply 70% ethanol with a cotton swab to sterilize the incision site.
   4. Remove sutures using fine tweezers and ophthalmic scissors. Irrigate surgery site and craniectomy with abundant sterile saline.
      1. Consider the animal for exclusion if the cortex displays disqualifying characteristics including excessive herniation, discoloration, disrupted vascularization, or hemorrhage.
   5. Load the cell transplant syringe
      1. Move cell suspension from the 37 °C incubator to a cell culture biosafety hood. Gently swirl or tap the tube to ensure a homogeneous cell suspension.
      2. Use a micro pipettor to load ~7.5 µL cell suspension into the Hamilton syringe through the plunger end.  
         1. Hold the syringe at a ~120° angle with the plunger end facing down. Insert the plunger, taking care not to introduce an air bubble between the suspension and plunger tip.
      3. Attach the gasket assembly to the pipette needle, then attach the needle to the syringe.
      4. Push the plunger to move cell suspension into the pipette needle.  If there is resistance against suspension outflow, use fine tweezers to break the needle tip to enlarge the diameter.
   6. Attach the syringe to the stereotaxic syringe pump.  Advance the plunger to make sure the syringe pump assembly is working properly.  
   7. Move the needle into the coordinates for injection.
      1. Align the needle tip to bregma. Set the X and Y coordinates to 0. Then move the needle tip over the craniectomy to 2.0 mm lateral and -1.0 mm posterior to bregma.  Touch the needle tip to the dura mater surface and set the stereotaxic coordinate to Z = 0. 
      2. Push the plunger to ensure the cell suspension is flowing adequately before introducing the needle into the brain.
      3. Introduce the needle into the brain to a depth of Z = -1.4 mm. These stereotaxic coordinates place the graft at the gray matter-white matter border of the deep cortex²⁰. 
   8. Start the syringe pump to infuse cell suspension. Set the lab bench timer to 15 min and start the timer. Use a long working distance microscope to monitor cell suspension outflow.
      1. Irrigate the surgery site with sterile saline during injection to maintain tissue hydration.
      2. At 15 min, slowly withdraw the transplantation needle.  Irrigate the surgery site with saline and close the incision with sutures.  
   9. Perform postoperative care
      1. Discontinue anesthesia. Deliver CsA by subcutaneous injection into scruff at 10 mg/kg dose. Place the mouse in a clean and pre-warmed postoperative cage.
      2. Provide acetaminophen analgesic in the drinking water at 1.0 mg/mL.
         NOTE: Provide analgesia according to the appropriate IACUC standard operating protocol (SOP) and in consideration of experimental outcome variables.
      3. Provide moistened chow food recovery cage to aid in rehydration and recovery.
6. Continue daily CsA injections at 10 mg/kg throughout the survival duration of the mouse.

3. Adhesive tape removal test of sensorimotor integration

1. Cut the electrical tape into 3 mm x 5 mm strips using a small razor knife prior to performing the behavior test. Use a smooth glass surface for cutting the adhesive strips.
   NOTE: Use yellow and red tape, as mice have difficulty distinguishing between these colors²¹.
   1. Select a small mirror that fits well inside the clear plastic box.  
      1. Fix the mirror in place at a roughly 45° angle with modeling clay or adhesive tape in order to view animal behavior from below.  
   2. Place the box and mirror assembly on a bench in a quiet dedicated behavior testing room.  Arrange the cylinder on the plastic box above the mirror.
   3. Arrange the handling cloth, tweezers, and adhesive strips on the bench next to the behavior testing box.
   4. Clean the box and cylinder with 70% ethanol and paper towels.  Allow surfaces to dry thoroughly.
   5. Bring the mice into the behavior testing room. Allow the mice to acclimate to the testing room for at least 30 min prior to behavior testing.  
   6. Remove water supplies from the cages to minimize urination events during testing, which can interfere with efficient test performance.
2. Perform the adhesive removal test
   NOTE: This behavior test is best performed by two to three investigators: one to operate the stopwatches, and one or two to handle the mice and observe the behavior.
   1. Use each tweezer to peel one adhesive strip of each color.  
      1. Choose which strip color corresponds to which forepaw and remain consistent throughout the trial.
   2. Use the handling cloth to restrain a mouse by the scruff of the neck and back such that the mouse holds the forepaws away from its body and head.
   3. Use tweezers to place an adhesive strip on the plantar surface of each forepaw. Use delicate and consistent finger pressure to secure the strips to the paws.
   4. Quickly place the mouse into the plastic cylinder. Start the two stopwatches when the mouse has all four paws on the plastic box.
   5. Use the two stopwatches to record the latencies for the following four events: left paw notice, left paw removal, right paw notice, right paw removal.
      1. Record a notice event when the animal makes unambiguous recognition of the adhesive strip by shaking or flinching the paw or biting the strip.
      2. Record a remove event when the animal effectively removes the strip from the forepaw plantar surface.  
         NOTE: The remove event is not disqualified by strip readherence or if the strip clings to the lateral surface of the forepaw.
      3. Stop the timer at 120 s if the corresponding strip has not been removed. 
      4. Record the elapsed times for the four events on the data sheet.
   6. Perform a second trial on each mouse
      1. Allow a minimum of 5 min to elapse between trials for an individual mouse to reduce stress, which can interfere with efficient performance on the test.
      2. Clean the apparatus with paper towels between testing mice to remove waste when testing multiple mice from a single cage.  
         1. Clean the apparatus thoroughly with 70% ethanol and paper towels when testing mice from multiple cages.
      3. Reverse the color-paw placement order for the second trial of the session to randomize any effect of color.
   7. Calculate the average latency of each event between two trials per daily session for each animal.
3. Clean the apparatus and handling cloth thoroughly with water, replace the cage water supplies, and return the mice to their holding facility.
4. Repeat steps 3.1-3.2 on successive experimental timepoints for a repeated measures trial design.
   NOTE: Repeat steps 3.2.1-3.2.5.3 daily for 5 days prior to any data collection to acclimate the animals to the task. Baseline data acquisition prior to surgery is recommended.

4. Diaminobenzidine (DAB) immunohistochemical analysis of graft survival and injury pathology

1. Euthanize the animals and perform a transcardial perfusion.
   1. Prepare solutions of 0.1 M phosphate-buffered saline (PBS) and 4% paraformaldehyde (PFA) in PBS. Balance the pH of both solutions to 7.4.
   2. Place the two solutions on ice in a fume hood. Place a peristaltic pump in the fume hood, and run the pump to fill the tubing with PBS. Set the pump flow rate to ~7 ml/min.  Connect a 25 G needle to the outflow tube of the pump.
   3. Place a drainage tray with a soft substrate in the hood. Raise one end of the tray at a slight angle so that perfusion fluids drain to the lower end.
   4. Place a mouse in an anesthesia induction chamber. Fill the chamber with 4% isoflurane using compressed 100% oxygen vehicle gas.
   5. Move the anesthetized mouse from the chamber to an anesthesia nose cone. Decrease the isoflurane to 2%.
   6. Perform a thoracotomy to expose the heart. Discontinue anesthesia, as the thoracotomy causes euthanasia.
   7. Carefully place the perfusion pump outflow needle in the left ventricle along the long axis of the heart. Do not pierce the heart septum, as this will cause poor perfusion.
   8. Use small scissors to cut the right atrium, then immediately turn on the peristaltic pump.
   9. Continue the PBS flow until the fluid leaving the right atrium runs clear of blood.
      1. Turn off the pump. Move the pump intake tube to the container of PFA. Restart the pump. Continue perfusing PFA until either the animal is sufficiently rigid or until a 20 mL volume has been pumped through.
      2. Turn off the pump. Remove the outflow needle from the heart. Move the intake tube from the PFA to the PBS, and thoroughly flush out PFA from the tubing.
2. Remove the brain from the skull by careful dissection. Place the brain in a small container filled with 4% paraformaldehyde, and allow the brain to post-fix overnight at 4 °C.
3. On the day after post-fixation, replace the P with PBS and 0.1% sodium azide. Store the brains in this solution until preparation for histologic sectioning.
4. Collect 40-50 µm sections of formaldehyde-fixed brain tissue via a freezing microtome or room temperature vibratome.
5. Pretreat tissues in 0.3% hydrogen peroxide in water to inactivate endogenous peroxidases, which can produce nonspecific staining.
6. Perform antigen retrieval using citric acid buffer (10 mM sodium citrate, 0.05% polysorbate-20, pH 6.0) at 60 °C for 30 min.
7. Perform antibody labeling by standard methods.  Refer to Lundell et al.’s report²² from this laboratory for more details.
   1. Use a mouse IgG neutralization kit (see Table of Materials) to reduce cross-reactivity with endogenous antibodies. Incubate the tissues in IgG neutralization buffer for at least 1 h at room temperature (RT), following the manufacturer’s directions.
   2. Use a mouse anti-human nuclear antigen primary antibody (hNA; 1:500 dilution) when transplanting human iPSC-derived cells to locate the transplanted cells. Incubate tissues with the primary antibody at 4 °C for 48-72 h using gentle agitation.
   3. After rinsing away primary antibody solution, incubate tissues with HRP-conjugated anti-mouse secondary antibody at 1:250 dilution for 2 h at RT.
   4. Perform DAB chromogen reaction by standard methods to reveal immunolabeling. Allow DAB reaction to develop for 5 to 10 min at RT.
8. Attach stained tissues to slides and apply cover slips using standard methods.
9. Quantify numbers of labeled cells using unbiased stereology as described previously^23,24. Perform analysis using 20 µm optical sections and a between section interval of three.