Induction of Intestinal Graft-versus-host Disease and Its Mini-endoscopic Assessment in Live Mice

The experimental methods described here have been approved by the government of Mittelfranken, Bavaria, Germany.

1. GvHD induction

1. Day 0: Total body irradiation of the recipient mice
   1. Use female CD45.2⁺ H2kd⁺ BALB/c mice that are at least 10 weeks old as recipients.
   2. Weigh and record the weight of the recipient mice prior to GvHD induction.
      NOTE: Ensure that the recipient has a minimal body weight of 20 g. The starting body weight will serve as the reference value to calculate the weight loss of the individual mouse over the course of GvHD induction and progression.
   3. Place up to five mice in the container of the X-ray irradiator.
   4. Irradiate the mice by total body irradiation with a single dose of 8 Gy of X-ray. Use Cs¹³⁷ as a radiation source.
2. Day 1: Reconstitution of the irradiated mice with T-cell-depleted bone marrow
   NOTE: The transplantation of allogeneic bone marrow cells, as outlined and detailed under section 1.2, should take place within 24 h after irradiation. Perform the procedures described below in a sterile tissue culture hood and use filtered reagents. Use allogeneic CD45.1/Ly5.1 B6.SJL-Ptprca Pepcb/BoyCrl mice as donors for T-cell-depleted bone marrow.
   1. Euthanize CD45.1/Ly5.1 B6.SJL mice that will donate allogeneic bone marrow cells in accordance with the institutional guidelines, 12-24 h after the irradiation of the recipient BALB/c mice. For this, anesthetize the CD45.1/Ly5.1 B6.SJL mice by inhalation of 5% isoflurane. Continue with the isoflurane exposure over 1 min after breathing arrest and confirm euthanasia by cervical dislocation.
   2. Disinfect the fur and skin of the mouse thoroughly with 70% ethanol.
   3. Position the mouse onto a clean working sheath so that the mouse is in a prone position, with its hindquarters facing the experimenter. Lift the fur at the Achilles' heel with the tip of a Semken forceps with a length of 13 cm and serrated curved tips. Incise the skin between the forceps and the heel with hardened 8.5 cm fine scissors with straight tips.
   4. Elongate the incision cranially (i.e., from the heel over the lower leg and thigh to the hip region). Remove the skin and fur from the hind limb with the help of the forceps.
   5. Perform the same procedure on the other hind limb.
   6. Remove the hind limbs by cutting through the adjacent hip joints.
   7. Cut away the rear paws. Cut through the knee joint. Store the thigh and shank of every hind limb together in a 92 mm Petri dish filled with phosphate-buffered saline (PBS) solution on ice.
   8. Clean the femur and tibia bones carefully by removing as much muscle tissue as possible from every thigh and shank. Transfer the femur and tibia bones to a 50 mL tube filled with sterile RPMI 1640 medium and place it on wet ice until all tibia and femur bones collected in step 1.2.7 are cleaned from muscle tissue.
   9. To isolate the bone marrow, transfer one femur or tibia bone at a time (which has been cleaned as described in step 1.2.8) from the 50 mL tube to a Petri dish (with a diameter of 92 mm) filled with RPMI 1640 medium. Cut off the ends of the femur or tibia with a scalpel to get access to the cavity of the bone containing the bone marrow.
   10. Prepare a 50 mL collection tube with 5 mL of RPMI 1640 medium. Insert a 26 G needle attached to a 1 mL syringe filled with 1 mL of RPMI 1640 medium into the bone cavity and flush the bone marrow out of the cavity into the collection tube by pushing the plunger.
   11. Repeat step 1.2.10 until the bone appears light and shiny (i.e., the bone cavity is visibly devoid of reddish bone marrow). Discard the empty bone.
   12. Repeat steps 1.2.9 – 1.2.11, using all femur and tibia bones from step 1.2.8, and collect all recovered bone marrow pieces in the same 50 mL collection tube from step 1.2.10. Keep the collection tube on wet ice until all bones from step 1.2.8 have been processed accordingly.
   13. Create a single-cell suspension by softly pipetting the flushed, crumbly bone marrow pieces up and down. Filter the bone marrow single-cell suspension through a 40 µm mesh screen cell strainer placed on a new 50 mL collection tube.
   14. Centrifuge the 50 mL collection tube containing the bone marrow single-cell suspension at 450 x g for 5 min at 4 °C. Discard the supernatant.
   15. Resuspend the pellet in 5 mL of ACK lysing buffer (1 mM Na₂EDTA; 10 mM KHCO₃; 144 mM NH₄Cl; pH 7.2) for red blood cell lysis. Incubate the cell suspension for 3 min at room temperature. Afterward, immediately add 10 mL of PBS solution to the cell suspension.
   16. Centrifuge the suspension at 450 x g for 5 min at 4 °C. Discard the supernatant and resuspend the cells in 2 mL of PBS solution.
   17. Count the bone marrow cells, using a hemocytometer. Preserve an aliquot of 6 x 10⁶ cells for flow cytometry analysis to check the purity of the cells after cell purification (see step 1.2.20).
   18. For the depletion of T cells from the bone marrow single-cell suspension, use a commercially available cell purification kit. Magnetically deplete CD90.2⁺ cells (i.e., lymphocytes) from the total bone marrow cells, following the manufacturer's protocol.
   19. Count the T-cell-depleted bone marrow cells that have been isolated as described in step 1.2.18, using a hemocytometer. Preserve an aliquot of 1 x 10⁶ cells for flow cytometry analysis to be performed later on, as described in step 1.2.20.
       NOTE: The on-average cell yield derived from one donor mouse is usually from 2 x 10⁷ to 3.4 x 10⁷ T-cell-depleted bone marrow cells.
   20. Confirm a successful T-cell depletion by flow cytometry. Stain 1 x 10⁶ cells, preserved from steps 1.2.17 (bone marrow cells before magnetic cell separation) and 1.2.19 (T-cell-depleted bone marrow cells after magnetic cell separation), with the following antibodies: α-CD45.1 (A20), α-CD3 (17A2), α-CD4 (GK1.5), and α-CD8α (53-6.7). Use the remaining cells from step 1.2.17 as an unstained control and for single-staining controls, to set up the flow cytometer.
       1. Transfer 1 x 10⁶ cells from each sample to a well of a 96-well plate with a V-bottom to perform the flow cytometric staining procedure.
       2. Spin down the cells within the 96-well plate (step 1.2.20.1) at 450 x g for 5 min at 4 °C. Discard the supernatant and resuspend the cells in 100 µL of FACS buffer (PBS supplemented with 3% filtered bovine serum).
       3. Centrifuge the cells within the 96-well plate at 450 x g for 5 min at 4 °C. Discard the supernatant.
       4. Add an appropriate amount of the antibody of choice (compare with step 1.2.20) to the samples for single staining in 100 µL of FACS buffer.
       5. Prepare a mixture of all antibodies (master mix) containing the appropriate amounts sufficient to separately stain the bone marrow cell aliquots preserved from prior (see step 1.2.17) and after (see step 1.2.19) magnetic T-cell depletion. Add 100 µL of the master mix to both aliquots.
       6. Add only 100 µL of FACS buffer to the unstained sample.
       7. Incubate the cells within the 96-well plate for 20 min at 4 °C in the dark.
       8. Add 100 µL of FACS buffer and centrifuge the cells within the 96-well plate at 450 x g for 5 min at 4 °C. Discard the supernatant and resuspend the cells in 100 µL of FACS buffer.
       9. Centrifuge the cells at 450 x g for 5 min at 4 °C. Discard the supernatant and resuspend the cells in 250 µL of FACS buffer.
       10. Transfer the samples to a 5 mL polystyrene round-bottom tube and analyze them with a flow cytometer instrument that is able to detect the fluorescent molecules that were used to label the antibodies employed to characterize the cell samples (see step 1.2.20).
       11. Compare the cell composition from before and after the magnetic cell separation.
           NOTE: A purity of approximately 95% CD45.1⁺CD3^– (i.e., T-cell-depleted bone marrow) cells within the live gate is usually achieved.
   21. Wash the T-cell-depleted bone marrow cell suspensions 2x with PBS solution (450 x g for 5 min at 4 °C) and, finally, resuspend the cells in PBS solution, adjusting the cell concentration to 5 x 10⁷ cells/mL. Keep the cells on wet ice until the injection.
   22. Inject T-cell-depleted bone marrow cells into the recipient mice, which were irradiated the day before (see section 1.1).
       1. Place the experimental mouse in a leak-proof chamber and induce anesthesia by inhalation of up to 4% isoflurane, until the mouse is unconscious. Confirm analgesia and loss of consciousness by testing the loss of the righting reflex and the subsequent loss of the pedal withdraw reflex.
       2. Inject 5 x 10⁶ T-cell-depleted bone marrow cells (i.e., 100 µL of 5×10⁷ cells/mL) containing PBS solution using a 1mL syringe equipped with a 30 G needle intravenously into the retrobulbar space containing the venous sinus.
3. Day 2: Transfer of T cells
   NOTE: Perform the described procedures below in a sterile tissue culture hood and use filtered reagents. Use CD45.2 C57Bl/6 (wild-type; WT) mice as donors for alloreactive T cells. The usage of congenic marker systems allows the distinguishability between recipient (CD45.2⁺ H2kd⁺ Balb/c mice) and donor hematopoietic cell types (bone marrow cells: CD45.1⁺ H2kb⁺ B6.SJL mice; allogeneic T cells: CD45.2⁺ H2kb⁺ C57/Bl6 mice).
   1. Euthanize C57Bl/6 mice in accordance with the applying institutional guidelines and authorities. Therefore, anesthetize mice by inhalation with a concentration of 5% isoflurane. Continue the isoflurane exposure until 1 min after breathing arrest and confirm euthanasia by cervical dislocation. Disinfect the fur and skin of the mouse thoroughly with 70% ethanol.
   2. Place a strainer with a 40 µm mesh screen on a 50 mL collection tube. Remove the spleen and place it onto the strainer.
   3. With a syringe plunger, comminute the spleen in the strainer. Wash the strainer and syringe plunger with PBS to collect all splenocytes.
   4. Centrifuge the cells within the 50 mL collection tube at 450 x g for 5 min at 4 °C. Discard the supernatant.
   5. Resuspend the splenocytes in 3 mL of ACK lysing buffer to lyse red blood cells. Incubate the cell suspension for 3 min. Afterward, add 10 mL of PBS and centrifuge the cells at 450 x g for 5 min at 4 °C. Discard the supernatant and resuspend the splenocytes in PBS.
   6. Count the spleen cells, using a hemocytometer. Divert an aliquot of 6 x 10⁶ cells for flow cytometry analysis, to evaluate the magnitude of purification in step 1.3.9.
   7. For a total CD3⁺ T-cell isolation from total splenocytes, use a commercially available cell purification kit. Isolate the splenic T cells, following the manufacturer's protocol.
   8. Count the splenic CD3⁺ T cells isolated as described in step 1.3.7, using a hemocytometer. Preserve an aliquot of 1 x 10⁶ T cells for flow cytometry analysis.
      NOTE: The on-average yield of splenic T cells per donor mouse isolated by magnetic cell separation ranges from 1.3 x 10⁷ to 2 x 10⁷ cells.
   9. Confirm the success of the T-cell isolation by flow cytometry. For a detailed description of the staining protocol, confer and follow steps 1.2.20.1 - 1.2.20.10.
      1. Stain 1 x 10⁶ splenocytes of steps 1.3.6 (before the magnetic cell separation) and 1.3.8 (after magnetic cell separation) with the following antibodies: α-CD45.2 (104), α-CD3 (17A2), α-CD4 (GK1.5), and α-CD8α (53-6.7).
      2. Use the remaining cells from step 1.3.6 as an unstained control and for single staining with the respective antibodies, to set up the flow cytometer.
      3. Compare the cell composition before and after magnetic cell separation.
         NOTE: A purity of ≥95% of CD45.2⁺CD3⁺ T cells within the lymphocyte live gate should be accomplished.
   10. Wash the T cells 2x with PBS (450 x g for 5 min at 4 °C) and, finally, resuspend the cells in PBS solution, adjusting the cell concentration to 7 x 10⁶ cells/mL. Keep the cells on ice until the injection.
   11. Inject alloreactive, splenic T cells (see step 1.3.10) into irradiated BALB/c mice who have previously received CD45.1⁺ T-cell-depleted bone marrow cells (see step 1.2.22).
       1. Induce anesthesia by placing the experimental mouse in a leak-proof chamber in which it is exposed to up to 4% isoflurane until it loses its consciousness. Confirm analgesia and loss of consciousness by testing the loss of the righting reflex and the subsequent loss of the pedal withdraw reflex.
       2. Inject 0.7 x 10⁶ alloreactive CD3⁺ T cells using a 1 mL syringe equipped with a 30 G needle  (i.e., 100 µL of 7 x 10⁶ cells/mL) containing PBS solution (see step 1.3.10), intravenously into the retrobulbar space of the mice, to induce GvHD. Denominate these mice as the experimental group.
       3. Inject a different group of mice with 100 µL of PBS alone, intravenously into the retrobulbar space, and denominate this group as the control group, subsequently called "no-T-cell-transplanted (noT) mice".

2. Assessment of Systemic GvHD

NOTE: Perform this procedure in a semi-sterile surrounding in an experimental room licensed and equipped for handling live mice within the animal facility.

1. Follow the clinical course of GvHD in individual mice; specifically, assess and score the experimental mice 3x a week for the presence of clinical GvHD symptoms, based on a scoring system described below and adapted from a scoring system established previously by Cook et al.⁶.
   1. Weigh each mouse individually, record the weight, and calculate thespecific body weight loss in reference to the body weight that was determined prior to the start of the experiment (corresponding to 100%) on day 0 (step 1.1.2). Score a weight loss of less than 10% of the starting weight with grade 0, a weight loss between 10% and 25% with grade 1, and a weight loss of more than 25% with grade 3.
   2. Score the activity level of each mouse, discriminating mice with a normal activity level (score = 0) from mice with a moderately decreased activity level (score = 1) and mice with stationary behavior unless they are externally stimulated (score = 2).
   3. Score the fur texture, thereby discriminating a normal, shiny, and kempt fur (score = 0) from a moderately ruffled fur (score = 1) and the presence of bald spots (score = 2).
   4. Score the skin texture, discriminating normal skin (score = 0) from sporadic spots of scaling skin (e.g., at the tail and the paws) (score = 1) and obvious scaling and sore skin areas (score = 2).
   5. Analyze the consistency of secreted stool fractions. Score stool with a normal (i.e., hard consistency) with grade 0, stool with deformable and soft stool with grade 1, and stool without any consistency and, hence, with severe diarrhea with grade 2.
   6. Sum up all separately scored parameters to a maximal clinical score of 12 per mouse.
2. Consider and evaluate the criteria of ceasing the experiment due to the current disease severity level of the individual mouse, in accordance with the applying institutional guidelines and authorized animal protocol.

3. Assessment of Intestinal GvHD

1. Scoring of GvHD-related lesions by mini-endoscopy of the distal colorectal region
   NOTE: Perform this in a semisterile surrounding in an experimental room licensed and equipped for handling live mice within the animal facility. Use a mini-endoscopic workstation that is approved for the use of small animals.
   1. Prepare the endoscopic device by covering the telescope (with a diameter of 1.9 mm and a length of 10 cm) with the endoscopic examination sheath. Clean and sterilize the endoscope with water and ethanol.
   2. Switch on the computer, the adjustable xenon light source, and the camera module, and set the focus in a way that objects close to the lens give a clear image.
   3. Connect the air pump to the endoscopic sheath. To visualize the air stream, dip the tip of the endoscope into a beaker glass filled with water. Regulate the air stream by adjusting the valve between the air pump and the endoscopic sheath. Adjust it to a slow, constant air stream, reflected by a few continuously ascending bubbles.
   4. Place a mouse into a leak-proof chamber. Induce anesthesia by inhalation of up to 4% isoflurane until the mouse is unconscious. Confirm analgesia and an absent state of the reflexes by testing the loss of the righting reflex and the subsequent loss of the pedal withdraw reflex.
   5. Use an appropriate veterinary anesthesia machine in accordance with the animal protocol used. Transfer the mouse from the chamber to the colonoscopy workspace. Here, position the anesthetized mouse, with its nose in the nosecone (which is connected to the anesthesia instrument), onto a clean working sheath so that the mouse is in a prone position, facing the experimenter with its hindquarters.
   6. To maintain anesthesia during the colonoscopy procedure, reduce the isoflurane concentration to approximately 2%. Monitor the mouse's respiration and response to stimulation during the colonoscopy and adjust the isoflurane concentration at the vaporizer if needed. Cover the mouse's eyes with salve to prevent them from getting dry.
   7. Control the efficacy of the anesthesia by pinching between the toes of the mouse. In the case of absent reactivity, proceed to step 3.1.8.
   8. Lift the tail just above the tail root with one hand and carefully insert the endoscope, positioned in the other hand, via the anus into the rectum.
   9. While the air stream inflates the colorectal lumen, slowly and carefully move the endoscope forward in aboral direction.
   10. Place the endoscope in the middle of the colonic lumen and maintain this position in order to minimize contact with the colonic wall and to avoid making scratches, deeper wall-related injuries (e.g., resulting in bleeding), or even wall perforation (see step 3.1.17). Control this step by permanently watching the video stream (i.e., under constant visualization of the colonic compartment).
   11. If feces constrict the view, remove the endoscope and perform an enema by rectally flushing with up to 2 mL of saline, using a soft Pasteur pipette. Use as little liquid as possible, to prevent an unintended dilution of the feces and other secondary effects negatively affecting the characteristics of the mucosal surface and, ultimately, the scoring results.
   12. Try to regularly position the endoscope at a defined (i.e., distinct) anatomic site within the distal colon to standardize the scoring of colonic inflammation and to optimize the comparability of scoring results between mice and across experiments.
       NOTE: Normally, with the rigid endoscope, a maximal depth of 4 cm can be reached via the rectal route. Between 2 and 4 cm aborally, the colon regularly turns into a flexure that cannot be passed with the rigid colonoscope. Use the colon region distally adjacent to the flexure as the default scoring spot.
   13. Start to record the video stream and begin with the scoring of inflammation.
   14. Evaluate the GvHD-related inflammation of the colon by scoring the parameters explained and depicted in Table 1 and Figure 3.
       1. Move the endoscope at the scoring spot gently and slightly back- and forward to assess the different parameters.
       2. Assess the parameter "translucency", as well as the "stool consistency", by positioning the endoscope in a wider distance in relation to the colonic wall.
       3. Assess the parameters "granularity" and "vascularity" by positioning the endoscope in close proximity to the colonic wall. For this task, carefully apply well-dosed tension to the colonic wall with the tip of the endoscope.
   15. Upon completion of the scoring process, stop recording.
       NOTE: Afterward, the recorded video clip can be used in total or used to generate representative images (i.e., screenshots) of the mini-endoscopically assessed criteria overall contributing to colonic inflammation.
   16. Carefully remove the endoscope.
   17. Discontinue isoflurane exposition by transferring the mouse to a separate cage in which it is exposed to ambient air. Warm the mouse with a red-light lamp and observe the mouse until it recovers from the anesthetic and regains full consciousness.
       NOTE: Due to the air inflation of the colon during the endoscopy, the abdominal region of the mouse may appear puffed up directly afterward. While this is normal and of transient nature, prolonged signs of massive and even progressive inflation of the abdomen and failure of recovery of the mouse might indicate an unintended perforation of the colon (in this case, the mouse needs to be euthanized immediately).
   18. Upon complete recovery, place the mouse back into its respective cage.
   19. Optional approach: It is also possible to take view-guided tissue biopsies. Take the following steps.
       NOTE: The colonoscopy will be performed in the same way as described in steps 3.1.1 – 3.1.18, with the following modifications.
       1. In step 3.1.1, use an endoscopic examination sheath with a built-in working channel, instead of a simple examination sheath without working channels, for covering the telescope. Introduce a biopsy forceps into the working channel until its tip is just visible in front of the endoscope at the video screen because this largely prevents unintended injury to the intestine.
       2. Proceed with steps 3.1.2 – 3.1.11. Insert the endoscope into the colon and push it carefully forward to the spot of interest.
       3. Employ the help of a second experimenter for the task of taking colonic tissue biopsies. Ask the second experimenter to navigate the tip of the biopsy forceps to the colonic region of choice by pushing the forceps within the working channel slowly forward until the jaws of the forceps can be opened. Constantly watch the video stream during this procedure to minimize the risk of injuring the colonic wall.
       4. Recover a colonic wall tissue sample by carefully opening and closing the jaws of the biopsy forceps.
          NOTE: Do this with caution to prevent the perforation of the colonic wall. In the rare case of a colonic perforation, immediately sacrifice the affected mouse by applying measurements in accordance with the institutional guidelines and under the continuous administration of anesthetics.
       5. Pull back and remove the closed forceps out of the working channel. Remove the specimen from the jaws of the biopsy forceps by dipping and carefully shaking the opened forceps in sterile PBS solution. Afterward, choose proper storage conditions for the tissue sample, in accordance with the planned downstream analyses. After disinfection with 70% ethanol, the forceps might be reused to take further biopsies.
       6. After taking biopsies, remove the endoscope and finish the colonoscopy as described in steps 3.1.16 – 3.1.18.
2. Histopathological analysis
   1. Between days 26 and 30 after the X-ray irradiation, euthanize the allo-HCT recipient mice in accordance with applying institutional guidelines and authorities. For this, anesthetize the mice by inhalation of 5% isoflurane. Continue with the isoflurane exposure for 1 min after breathing arrest and confirm euthanasia by cervical dislocation. Thoroughly disinfect the fur and skin of the mouse with 70% ethanol.
   2. Open the abdominal cavity and remove the colon. Transfer it to a Petri dish (with a diameter of 92 mm) with PBS.
   3. Fill a 5 mL dispenser tip with PBS. With the help of a Semken forceps with a length of 13 cm and serrated curved tips, slip the distal part of the colon (approximately 5 mm) over the tip of the dispenser tip. Carefully fix the colon with the forceps at the dispenser tip and flush the colon with PBS by pressing down the plunger of the dispenser tip to remove feces from the intestine.
   4. Cut out a colonic segment located about 5 mm from the anus, with the help of a scalpel.
   5. Fix the resected gut specimen in 4.5% formaldehyde overnight. Before embedding it in paraffin, place the tissue in an upright position.
   6. Cut 3 µm cross sections of paraffin-embedded colon tissue and stain them with hematoxylin and eosin (HE), using standard staining protocols.
   7. Prepare HE-stained cross sections of colon tissue samples.
   8. Adapted from the scoring matrix reported by Kaplan et al.¹², histopathologically score the inflammatory activity semiquantitatively, with scores from 0-3: no inflammation (score = 0), mild inflammation (score = 1), moderate inflammation (score = 2), and severe inflammation (score = 3).
      NOTE: Ideally, employ for this task the expertise of a pathologist who is experienced in the evaluation of murine intestinal GvHD and blinded to the nature of the experimental and control groups.