Assessment of Chimeric Antigen Receptor T Cell-Associated Toxicities Using an Acute Lymphoblastic Leukemia Patient-Derived Xenograft Mouse Model

Podsumowanie

Here, we describe a protocol in which an acute lymphoblastic leukemia patient-derived xenograft model is used as a strategy to assess and monitor CD19-targeted chimeric antigen receptor T cell-associated toxicities.

Streszczenie

Chimeric antigen receptor T (CART) cell therapy has emerged as a powerful tool for the treatment of multiple types of CD19⁺ malignancies, which has led to the recent FDA approval of several CD19-targeted CART (CART19) cell therapies. However, CART cell therapy is associated with a unique set of toxicities that carry their own morbidity and mortality. This includes cytokine release syndrome (CRS) and neuroinflammation (NI). The use of preclinical mouse models has been crucial in the research and development of CART technology for assessing both CART efficacy and CART toxicity. The available preclinical models to test this adoptive cellular immunotherapy include syngeneic, xenograft, transgenic, and humanized mouse models. There is no single model that seamlessly mirrors the human immune system, and each model has strengths and weaknesses. This methods paper aims to describe a patient-derived xenograft model using leukemic blasts from patients with acute lymphoblastic leukemia as a strategy to assess CART19-associated toxicities, CRS, and NI. This model has been shown to recapitulate CART19-associated toxicities as well as therapeutic efficacy as seen in the clinic.

Wprowadzenie

Chimeric antigen receptor T (CART) cell therapy has revolutionized the field of cancer immunotherapy. It has proven to be successful in treating relapsed/refractory acute lymphoblastic leukemia (ALL), large B cell lymphoma, mantle cell lymphoma, follicular lymphoma, and multiple myeloma^{1,2,3,4,5,6,7}, leading to recent FDA approvals. Despite the initial success in clinical trials, treatment with CART cell therapy results in toxicities that are often severe and occasionally lethal. The most common toxicities after CART cell therapy include the development of CRS and NI, also referred to as immune effector cell-associated neurotoxicity syndrome (ICANS)^8,9. CRS is caused due to the overactivation and massive expansion of CART cells in vivo, leading to the subsequent secretion of multiple inflammatory cytokines, including interferon-γ, tumor necrosis factor-α, granulocyte-macrophage colony-stimulating factor (GM-CSF), and interleukin-6 (IL-6). This results in hypotension, high fevers, capillary leak syndrome, respiratory failure, multi-organ failure, and in some cases, death^10,11. CRS develops in 50-100% of cases after CART19 cell therapy^11,12,13. ICANS is another unique adverse event associated with CART cell therapy and is characterized by generalized cerebral edema, confusion, obtundation, aphasia, motor weakness, and occasionally, seizures^9,14. Any grade of ICANS occurs in up to 70% of patients, and Grades 3-4 are reported in 20-30% of patients^5,10,15,16. Overall, CRS and ICANS are common and can be fatal.

The management of ICANS after CART cell therapy is challenging. Most patients with ICANS also experience CRS¹⁷, which can often be treated with the IL-6 receptor antagonist tocilizumab or steroids¹⁸. A previous report revealed that early intervention with tocilizumab decreased the rate of severe CRS but did not affect the incidence or severity of ICANS¹⁹. Currently, there is no effective treatment or prophylactic agent for ICANS, and it is crucial to investigate preventive strategies²⁰.

Myeloid cells and associated cytokines/chemokines are thought to be the main drivers of the development of CRS and ICANS²¹. While CRS is directly related to the extreme elevation of cytokines and T cell expansion, the pathophysiology of ICANS is largely unknown^22,23. Therefore, it is imperative to establish a mouse model that recapitulates these toxicities after CART cell therapy to study the mechanisms and develop preventive strategies.

There are multiple preclinical animal models currently used to study, optimize, and validate the efficacy of CART cells, as well as to monitor their associated toxicities. These include syngeneic, xenograft, immunocompetent transgenic, humanized transgenic, and patient-derived xenograft mice, in addition to primate models. However, each of these models has drawbacks, and some do not reflect the true efficacy or safety concerns of CART cells^24,25. Therefore, it is imperative to carefully choose the best model for the intended goals of the study.

This article seeks to describe the methodology that is used to assess CART cell-associated toxicities, CRS and NI, using an ALL patient-derived xenograft (PDX) in vivo model (Figure 1). Specifically, in the methods described here, CART19 cells generated in the authors' laboratory are used following previously described protocols. Briefly, human T cells are isolated from healthy donor peripheral blood mononuclear cells (PBMCs) via a density gradient technique, stimulated with CD3/CD28 beads on day 0, and lentivirally transduced on day 1 with CARs composed of a CD19-targeted single chain variable fragment fused to 4-1BB and CD3ζ signaling domains. These CART cells are then expanded, de-beaded on day 6, and cryopreserved on day 8^{26,27,28,29,30}. As outlined previously, mice are subjected to lymphodepleting treatment, followed by the administration of patient-derived leukemic blasts (ALL)²⁸. First, tumor engraftment is verified via submandibular blood collection. Following the establishment of an appropriate tumor burden, CART19 cells are administered to the mice. Then, the mice are weighed daily to assess well-being. Small animal magnetic resonance imaging (MRI) is performed to assess NI, along with tail bleeding to assess T cell expansion and cytokine/chemokine production. The techniques described below are highly recommended to be used as a model to study CART cell-associated toxicities in a PDX model.

Protokół

This protocol follows the guidelines of Mayo Clinic's Institutional Review Board (IRB), Institutional Animal Care and Use Committee (IACUC A00001767), and Institutional Biosafety Committee (IBC, Bios00000006.04).

NOTE: All the materials used to work with mice must be sterile.

1. Injection of busulfan to NSG mice

1. Obtain male, 8-12 weeks old, immunocompromised, NOD-SCID IL2rγnull (NSG) mice, and weigh them prior to injection.
   NOTE: For statistical significance, it is recommended to use at least five mice per group and to repeat this experiment at least once more with female mice.
2. Prepare busulfan for intraperitoneal (i.p.) injection. Using a 1 mL insulin syringe, slowly and carefully fill the syringe with exactly 30 mg/kg of busulfan, followed by i.p injection to the mice, and place them back into the cage.

2. Injection of ALL patient-derived blasts (CD19⁺) to the NSG mice

NOTE: This protocol follows the guidelines of Mayo Clinic's Institutional Biosafety Committee (IBC, Bios00000006.04).

1. Collect primary leukemic blasts from the peripheral blood of patients with relapsed and refractory ALL.
2. Prepare target cells for intravenous (i.v.) injection.
   1. Count CD19⁺ target cells using a hemocytometer, and record the final volume and concentration of cells.
      NOTE: To determine cell viability, it is strongly recommended to utilize trypan blue while counting the cells.
   2. Pellet the cells by centrifugation at 300 × g for 5 min at 4 °C.
   3. Resuspend the cell pellet in phosphate-buffered saline (PBS) to a final concentration of 50 × 10⁶ cells/mL in a 1.5 mL microcentrifuge tube, and place on ice.
   4. Finger-vortex the 1.5 mL microcentrifuge tube containing the target cells until the cells are completely homogenized.
   5. Using a 1 mL insulin syringe, slowly and carefully fill the syringe with exactly 100 µL of the prepared target cells, and flick the syringe to remove bubbles.
      NOTE: The volume of 100 µL will administer 5 × 10⁶ cells to each mouse.
   6. Place the mice under warm light for 5-10 min. Once the tail vein becomes engorged and more visible to the naked eye, place the mice in an appropriate confinement/restraining chamber.
   7. Wipe the tail of the mouse with an alcohol swab. Inject the target cells directly into the tail vein (i.v.), and place the mouse back in the cage. After the inoculation of the target cells, monitor the mice daily or as recommended by the local animal ethics committee.

3. Tumor engraftment assessment

1. Starting at 6 weeks after the target cell injection, assess tumor engraftment by using flow cytometry to measure the expression of CD19⁺ cells in the peripheral blood.
2. Peripheral blood assay to assess CD19⁺ expansion
   1. Withdraw blood from the mice 6-8 weeks after CART19 cell injection.
      NOTE: Several methods can be used to withdraw blood. For this protocol, submandibular blood collection is the preferred method of choice.
   2. Anesthetize the mouse with isoflurane in an induction chamber with 5% isoflurane. Once anesthesia is achieved, maintain at 1-3% isoflurane inhaled through a nose cone.
      NOTE: It is strongly recommended to apply ophthalmic ointment to prevent eye dryness.
   3. Remove the mouse from the chamber, and hold the mouse with the non-dominant hand by grasping the loose skin over the neck. Puncture the vein with a lancet slightly behind the mandible.
      NOTE: Only the tip of the needle (1-2 mm) is required to enter the vein.
   4. As the blood is dripping, collect at least 50 µL in a heparin-coated tube, and mix well by inverting the tube up and down several times. Transfer 30 µL of blood into a 5 mL round bottom polystyrene tube.
      NOTE: For the remaining blood, spin down the tube at 17,000 × g for 10 min at 4 °C. Transfer the serum (supernatant layer) to a clean and labeled 1.5 mL microcentrifuge tube, and store it at −80 °C (for use in the cytokine assay).
   5. Add 2 mL of lysis buffer (10x ammonium chloride-based lysing buffer diluted to 1x using nuclease-free water) into the 5 mL round bottom tube containing the blood. Mix very well by vortexing the tube. Incubate the tube at room temperature for 20 min to ensure proper lysis of the red blood cells.
   6. Add 2 mL of flow buffer (PBS, 0.2 g/L of potassium chloride, 0.2 g/L of potassium phosphate monobasic, 8 g/L of sodium chloride, and 1.15 g/L of sodium phosphate dibasic containing 2% fetal bovine serum and 1% sodium azide).
   7. Centrifuge the tube at 300 × g for 5 min at 4 °C. Decant the supernatant, and add 2 mL of flow buffer. Repeat this washing step 2x.
   8. Carefully decant the contents of the tube. Observe the small amount of fluid with a pellet remaining in the tube. Add 100 µL of flow buffer, and resuspend very well.
   9. Transfer all the remaining liquid in the 5 mL flow tube into a 96-well plate. Make sure each sample is properly labeled. Add 100 µL of flow buffer to the corresponding wells and centrifuge the 96-well plate at 650 × g for 3 min at 4 °C.
3. Assess the expansion of the CD19⁺ target cells by flow cytometry (Figure 2).
   1. Prepare a master mix containing 0.3 µL of live/dead stain, 0.13 µg (2.5 µL) of anti-human CD19 antibody, 0.06 µg (2.5 µL) of anti-human CD45 antibody, 5 µg (2.5 µL) of anti-mouse CD45, and 42.2 µL of flow buffer for a total of 50 µL per sample. Incubate in the dark at room temperature for 15 min.
   2. Wash with 150 µL of flow buffer followed by centrifugation at 650 × g for 3 min at 4 °C. Decant the supernatant, and resuspend the pellet in 195 µL of flow buffer and 5 µL of counting beads.
   3. Acquire on a flow cytometer to determine the level of CD19⁺ cell expression. For gating and analysis purposes, first gate the population of interest (CD19⁺) based on the forward versus side scatter characteristics, followed by a single gating population, and then live cells. Once the live cells are gated, assess the human CD45 versus mouse CD45 population, and from the human population, gate the CD19⁺ target population.
4. Quantify to determine the amount present in 70 µL, and then express in cells/µL using equation (1):
   Absolute CD19⁺ cell count = ([CD19⁺ cells/counting beads] × number of counting beads within 5 µL)/70
   ​​NOTE: Repeat the peripheral blood assay every 5 days to assess the CD19⁺ cell expansion. Collect no more than 200 μL of blood cumulatively every 2 weeks in accordance with local animal ethics committee guidelines. Once the mice have reached a disease burden of ≥10 cells/µL, randomize to CART19 cell or control groups (section 4).

4. Administration of CART19 cells  in vivo

1. Preparation of CART19 cells (~Day 80):
   NOTE: The generation of CART19 cells has been previously published^27,30,31. This procedure must be carried out inside a Biosafety Cabinet Level 2+ using aseptic technique and personal protective equipment.
   1. Thaw the CART19 cells in a water bath container (37 °C). Then, wash the cells in warm T cell medium (TCM) to remove the dimethyl sulfoxide.
      NOTE: TCM is made of 10% human AB serum(v/v), 1% penicillin-streptomycin-glutamine (v/v), and serum-free hematopoietic cell medium^27,30.
   2. Centrifuge the CART19 cells at 300 × g for 8 min at 4 °C, and resuspend in warm TCM to a final concentration of 2 × 10⁶ cells/mL. Allow the CART cells to rest in an incubator (37 °C, 5% CO₂) overnight but for no more than 16 h.
2. Administration of CART19 cells via i.v. injection (Day ~80)
   1. Count the CART19 cells, and spin down at 300 × g for 8 min at 4 °C.
   2. Resuspend the CART19 cells with 10 mL of PBS, and spin down at 300 × g for 8 min at 4 °C.
   3. Resuspend the CART19 cells to a final concentration of 40 × 10⁶ cells/mL with PBS, transfer the cells into a sterile 1.5 mL microcentrifuge tube, and place them on ice.
   4. Perform CART19 cell tail vein injections (as detailed above) by injecting 100 µL of resuspended cells i.v.
      ​NOTE: A volume of 100 µL will administer 4 × 10⁶ cells to each mouse. Include a group of untreated mice as a negative control.

5. Assessment of CART19 cell-associated toxicities

1. After CART19 cell administration, monitor the mice 2x a day to assess any changes in their well-being, such as motor weakness, hunched body, and loss of body weight.
   NOTE: All these physical changes correlate with CRS symptoms in this model²⁸.
2. Once the mice begin to develop motor weakness and weight reduction (10-20% reduction from the baseline), collect peripheral blood to analyze the tumor burden and conduct serum isolation for cytokines as per the methodology described in section 3 (Figure 3).
3. Store the serum microcentrifuge tubes at −80 °C, and use the sera to analyze the chemokines and cytokines using a Multiplex assay (Figure 4).
   NOTE: If the mice exhibit signs of hind limb paralysis, appear moribund or lethargic, and the weight loss is > 20% of their body weight up to the point that limits their ability to reach food and/or water, then they will be subjected to euthanasia.

6. MRI imaging

NOTE: A preclinical (for small animals) MRI scanner with a vertical bore magnet was used for in vivo magnetic resonance and image acquisition^32,33.

1. Mix 120 µL of Gadolinium + 880 µL of saline solution, and load into a 1 mL insulin syringe. Inject 100 µL into each mouse (i.p.).
   NOTE: Gadolinium should be injected 15-20 min prior to the start of the experiment.
2. Anesthetize the mouse using isoflurane (2-3%) in an induction chamber for 10-15 min.
   NOTE: It is highly recommended to apply ophthalmic ointment to prevent eye dryness.
3. Place the mouse spine on the rodent-compatible cradle probe. Then, proceed to position the mouse by hooking its teeth on the bite bar.
4. Pull the head of the mouse into the 25 mm volume coil, and adjust the nose cone tethered to the isoflurane anesthesia system. Tighten the knob of the bite bar to maintain the position for the duration of the scan.
   NOTE: If conducting a contrast-enhanced MRI scan, ensure that the gadolinium (in this case) or contrast agent is injected at least 10 min prior to placing the apparatus into the bore magnet. Throughout the time of the imaging, the mice will be anesthetized by the inhalation of 2-3% isoflurane in air via the nose cone, and their respiratory rate will be simultaneously monitored.
5. For breathing assessment, use a breathing/respiratory monitoring device.
   NOTE: It is important to assess the body temperature as well in order to prevent possible hypothermia due to extended exposure to anesthesia. It is recommended to use heat pads to control the body temperature.
   1. Attach the breathing monitor probe close to the diaphragm, and secure it with surgical tape. Keep the respiration rate between 20-60 breaths per min so that the mouse's condition is stable.
      NOTE: This rate provides more stable MRI images and prevents motion artifacts in the acquired image. If the respiratory rate is higher than the 20-60 breaths per min range, increase the concentration of isoflurane. If the respiratory rate is less than 20 breaths per min, the anesthesia is too deep; therefore, concentration needs to be decreased.
6. Insert the animal probe into the small bore within the vertical-bore small-animal MRI system. Adjust the animal's head at the center of the coil. Secure the apparatus with the lock so it can stand upright, and connect the instrument to the computer.
7. Open and use the software (e.g., Paravision) to set up the scan positions and types of scans. Determine the optimal axial and sagittal positions while keeping them consistent for all the experimental groups.
   NOTE: It is recommended to use the acquisition of a trial scan as a reference prior to the actual full-length MRI scans.
8. Proceed with the MRI data collection (as previously described^32,34,35,36). Obtain both T1-weighted and T2-weighted MRI images using the suggested setup mentioned below.
   1. For T1-weighted MRI images, use T1-weighted multislice multiecho (MSME) sequence with a repetition time (TR) of 300 ms, an echo time (TE) of 9.5 ms, a field of view (FOV) of 4 cm x 2 cm x 2 cm, and a matrix of 192 x 96 x 96.
   2. For T2-weighted MRI images, use a Multislice Multiecho (MSME) sequence with a TR of 300 ms, a TE of 9.5 ms, an FOV of 4 cm x 2 cm x 2 cm, and a matrix of 192 x 96 x 96 was used.
9. Once the scans are complete, remove the probe from the bore. Gently extract the mouse by removing the teeth off the bite bar. Monitor the animal in a separate cage until it is fully conscious and has regained adequate motor function.
10. After the extraction of data from the software, use analysis software for the quantification and 3D volume rendering (Figure 4) of the hyperintensity regions.
    NOTE: If possible, it is strongly encouraged to have two separate blinded observers for the data analysis.

Wyniki

The aim of this protocol is to assess CART cell-associated toxicities using a PDX mice model from tumor cells of patients with ALL (Figure 1). First, NSG mice received i.p. injections of busulfan (30 mg/kg) with the goal of immunosuppressing them and facilitating CART cell engraftment²⁸. The following day, they received ~5 × 10⁶ PBMCs (i.v.) derived from ALL patients. The mice were monitored for engraftment for ~13 weeks via the tail bleeding ...

Dyskusje

In this report, a methodology to assess CART cell-associated toxicities using an ALL PDX model has been described. More specifically, this model seeks to mimic two life-threatening toxicities, CRS and NI, that patients often experience after the infusion of CART cells. It recapitulates many hallmarks of CART toxicities observed in the clinic: weight loss, motor dysfunction, neuroinflammation, inflammatory cytokine and chemokine production, and the infiltration of different effector cells into the central nervous system

Materiały

Name	Company	Catalog Number	Comments
APC Anti-Human CD19	Biolegend	302211
Alcohol Prep Pad	Wecol	6818
Analyze 14.0 software	AnalyzeDirect Inc.	N/A	https://analyzedirect.com/analyze14/
Artificial tears (Mineral oil and petrolatum)	Akorn	17478-062-35	Topical ophtalmic gel to prevent eye dryness
BD FACS Lysing Solution	BD	349202	Red blood cells lysing buffer
BD Micro-Fin IV insulin syringes	BD	329461
Brillian Violet 421 Anti-Human CD45	Biolegend	304032
Bruker Avance II 7 Tesla	Bruker Biospin	N/A	MRI machine
Busulfan (NSC-750)	Selleckchem	S1692
CountBright absolute counting beads	Invitrogen	C36950
CytoFLEX System B4-R2-V2	Beckman Coulter	C10343	flow cytometer
Dulbecco's Phosphate-Buffered Saline	Gibco	14190-144
ERT Control/Gating Module	SA Instruments	Model 1030	Small Animal Monitoring Respiratory and Gating System
Fetal bovine serum	Millipore Sigma	F8067
Hemocytometer	Bright-Line	Z359629-1EA
Human AB Serum; Male Donors; type AB; US	Corning	35-060-CI
Isoflurane (Liquid)	Sigma-Aldrich	792632
LIVE/DEAD Fixable Aqua Dead Cell Stain Kit, for 405 nm excitation	Invitrogen	L34966
Microvette 500 Lithium heparin	Sarstedt	20.1345.100	Blood collection tube
MILLIPLEX Huma/Cytokine/Chemokine Magnetic Beads Panel	Millipore Sigma	HCYTMAG-60K-PX38	Immunology Multiplex Assay to identify cytokines and chemokines
Omniscan	Ge Healthcare Inc.	0407-0690-10	Gadolinium-based constrast agent
Pd Anti-Mouse CD45	Biolegend	103106
Penicillin-Streptomycin-Glutamine (100x), Liquid	Gibco	10378-016
Round Bottom Polysterene Test tube	Corning	352008
Sodium Azide, 5% (w/v)	Ricca Chemical	7144.8-16
Stainless Steel Surgical Blade	Bard-Parker	371215
X-VIVO 15 Serum-free Hematopoietic Cell Medium	Lonza	04-418Q