Waiting
Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove
JoVE Journal Cancer Research

Cancer Research

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

Published: February 10, 2023 doi: 10.3791/64535
Automatic Translation
1,2,3,4,5, 1,2, 1,2, 6, 6, 1,2, 1,2, 6, 1,2,6
1T Cell Engineering Laboratory, Mayo Clinic, Rochester, 2Division of Hematology, Mayo Clinic, Rochester, 3Mayo Clinic Graduate School of Biomedical Sciences, Mayo Clinic, Rochester, 4Department of Molecular Medicine, Mayo Clinic, Rochester, 5Regenerative Sciences PhD Program, Mayo Clinic, Rochester, 6Department of Immunology, Mayo Clinic, Rochester

ERRATUM NOTICE

Summary

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.

Abstract

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.

Introduction

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 myeloma1,2,3,4,5,6,7, 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, death10,11. CRS develops in 50-100% of cases after CART19 cell therapy11,12,13. 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, seizures9,14. Any grade of ICANS occurs in up to 70% of patients, and Grades 3-4 are reported in 20-30% of patients5,10,15,16. 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 unknown22,23. 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 cells24,25. 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 826,27,28,29,30. 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.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

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 published27,30,31. 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 medium27,30.
    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 acquisition32,33.

  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 described32,34,35,36). 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.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

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 assay. Red blood cells were collected and lysed, followed by flow cytometry preparation to assess the expression of CD19+ tumor cells (Figure 2).

Once the mice reached ≥10 tumor cells/µL, they were randomized and prepared to receive 4 × 106 CART19 cells (i.v.). To assess CART19-associated toxicities, the mice were weighed once per day (Figure 3A). Weight loss was chosen as a symptom of CRS appearance, as it has been previously revealed to be related to CART cell-associated toxicities28,37. Peripheral blood assays (via submandibular blood collection) were performed to assess tumor burden and CART cell expansion by flow cytometry analysis of human CD19+ tumor cells and human CD3+ T cells. In addition, serum was collected, and an assessment of inflammatory cytokines was performed. Assays were done one day prior to CART19 cell infusion as well as at 5 and 10 days after CART19 cell infusion, as these timepoints are particularly relevant to CRS onset and development of symptoms.

Here, in particular, a Multiplex assay was performed before and after CART19 cell administration, where the most representative cytokines and chemokines were GM-CSF, IL-18, MIP-1α, and IP-10 (Figure 3B). However, other assays can be done to assess the concentrations of these proteins in the serum. Alternatively, mice were subjected to MRI scanning to evaluate vascular permeability in the central nervous system (CNS). Mice were injected with the contrast reagent gadolinium (i.p.) to measure brain inflammation and blood-brain barrier disruption38. Then, they were anesthetized with isoflurane, and contrast-enhanced T1-weighted and T2-weighted images were taken (Figure 4). With T2 images (Figure 4, left), the brighter section (white) indicates the presence of edema or fluid, which is mostly seen in brain tumors and inflammation. With T1 images (Figure 4, middle), the brighter sections indicate areas where there is blood leaking or blood-brain barrier disruption. Lastly, with the help of software, the 3D reconstruction images (Figure 4, right) were assembled using gadolinium-enhanced T1 images based on hyperintensity signals corresponding to vascular permeability, which renders the volume of gadolinium leakage in the brain34.

Figure 1
Figure 1: Experimental scheme of a patient-derived xenograft model used to assess CART cell-associated toxicities, cytokine release syndrome, and neuroinflammation. NSG mice were conditioned with 30 mg/kg busulfan (i.p.) and received 3 × 106-5 × 106 blasts (i.v.) derived from the peripheral blood of patients with ALL. Next, the mice were monitored for tumor engraftment via submandibular blood collection, followed by flow cytometry to assess CD19+ blasts. When the peripheral blood CD19+ cells were ≥10 cells/µL, the mice received 2 × 106-5 × 106 CART19 cells. The mice were weighed daily as a measure of their well-being. At 10 days post CART19 cell infusion, mouse brain MRIs were performed to detect neuroinflammation. Tail bleeding to assess cytokine/chemokine production and T cell expansion was performed weekly post CART19 cell injection. Abbreviations: CART = chimeric antigen receptor T cell; CART19 = CD19-targeted CART; ALL = acute lymphoblastic leukemia; i.v. = intravenous; i.p. = intraperitoneal; PB = peripheral blood; NSG = NOD-SCID IL2rγnull. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Analysis of CD19+ tumor cells as an indicator of tumor burden in an ALL PDX model to assess CART-cell associated toxicities. (A) Representative flow cytometric analysis showing the gating strategy to analyze the CD19+ cell population taken from a peripheral blood sample of a mouse injected with patient-derived ALL blasts. (B) The quantification and comparison of the CD19+ cell populations between mice treated with CART19 versus untreated mice based on raw data obtained from flow cytometry analysis. (one-way ANOVA; n = 3 mice per group; error bars, SEM). Abbreviations: ns = not significant; ALL = acute lymphoblastic leukemia; PDX = patient-derived xenograft; CART = chimeric antigen receptor T cell; CART19 = CD19-targeted CART; SSC-H = side scatter-peak height; FSC-H = forward scatter-peak height; SSC-A = side scatter-peak area; FSC-A = forward scatter-peak area. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Assessment of CART19-associated toxicities using an ALL PDX model. (A) Representative graph of the percent weight reduction in mice treated with CART19 cells or control untransduced T cells for the first 6 days after CART (two-way ANOVA, ***p < 0.001, ****p < 0.00001; error bars, SEM). (B) Expression of cytokines and chemokines in NSG mice serum before and after CART19 cell administration. NSG mice were subjected to submandibular bleeding before and after CART19 cell administration (i.v.) to collect their serum and assess the expression of the following cytokines and chemokines: GM-CSF, IL-18, MIP-1α, IP-10, IL-1β and IL-6 (two-way ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001; n =3 mice; nine technical replicates; error bars, SEM). Abbreviations: ns = not significant; ALL = acute lymphoblastic leukemia; PDX = patient-derived xenograft; CART = chimeric antigen receptor T cell; CART19 = CD19-targeted CART; NSG = NOD-SCID IL2rγnull; GM-CSF = granulocyte-macrophage colony-stimulating factor; IL-18 = interleukin-18; MIP-1α = macrophage inflammatory protein-1 alpha; IP-10 = interferon gamma-induced protein 10; IL-1β = interleukin-1β; IL-6 = interleukin-6. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Magnetic resonance imaging used to assess neuroinflammation during CART19 cell treatment in a PDX model. NSG mice engrafted with tumor cells derived from ALL patients were infused with CART19 cells, and the brains were imaged with gadolinium-enhanced MRI after the mice exhibited neurological symptoms within 10 days after CART infusion. (Left) T2-weighted image revealing evidence of edema and possible inflammatory infiltrate during NI in CART19-treated mice (red arrow). (Middle) Representative axial section of T1-weighted MRI showing contrast enhancement within the brain parenchyma, indicating increased vascular permeability in CART19-treated mice (red arrow). (Right) Three-dimensional reconstructions of the rodent brain with T1-hyperintensity regions (red) were generated using Analyze 12.0 software. Abbreviations: MRI = magnetic resonance imaging; ALL = acute lymphoblastic leukemia; PDX = patient-derived xenograft; CART = chimeric antigen receptor T cell; CART19 = CD19-targeted CART. Please click here to view a larger version of this figure.

Human Mouse
Cytokine/Chemokine Human myeloid cytokines (IL-6, GM-CSF, IL-1, MCP-1) Human myeloid cytokines (IL-6, GM-CSF, IL-1, MCP-1)
Time to CRS (Day) 2-5 4-6
Time to NI (Day) 5-7 6-8
BBB status BBB disruption BBB disruption
Tissue resident macrophage Microglial activation Microglial activation
CART cell in CNS Brain infiltration Brain infiltration

Table 1: Recapitulation of the clinically observed CART cell-associated toxicities by the ALL PDX model. Abbreviations: GM-CSF = granulocyte-macrophage colony stimulating factor; IL = interleukin; CRS = cytokine release syndrome; NI = neuroinflammation; BBB = blood-brain barrier; CNS = central nervous system; MCP-1 = monocyte chemoattractant protein 1.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

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 system8,20,28. Previously, this PDX model has been used to mimic the initial development of CRS and NI (Table 1) and assess different strategies for toxicity prevention and treatment. One study in this PDX model utilized CART19 cells depleted of GM-CSF. More specifically, these results demonstrated that GM-CSF depletion resulted in the reduced secretion of CRS-linked cytokines and chemokines, which was clearly demonstrated using this ALL PDX mouse model29. This PDX model has also been used successfully to assess CART cell in vivo expansion correlating with CRS development after CART19 cell infusion using a sodium iodide symporter (NIS) as an imaging platform26. Therefore, this proposed model represents a useful tool that can be used to assess the potential toxicities arising from CART cells.

It is important to mention that many other models have been used to study CART cells in vivo. These include syngeneic, transgenic, and humanized mouse models, in addition to xenogeneic models24. The syngeneic mouse model, also called the immunocompetent allograft mouse model, utilizes CART cells, tumor, and target antigens that are murine in origin. This is advantageous as it allows for the study of CART cells in a fully functional immune system and can also reveal on-target, off-tumor toxicities. The disadvantage of this model is that mouse biology and immunity are different from those of humans, and, therefore, it is not a relevant model for translational studies where the goal is to proceed to clinical trials24,39.

Immunocompetent transgenic mouse models express a human tumor-associated antigen (TAA) transgene on both healthy tissue and tumors in patterns mimicking those found in human tissues. These mice have also been used in CART studies to better evaluate the safety of CART cells by revealing on-target off-tumor effects40,41. Humanized mice implanted with CD34+ human hematopoietic stem cells can be an advantageous model to study CART efficacy and toxicity on the human hematopoietic system, as the reconstituted immune system can mimic CART inhibition and induce a cytokine storm that recapitulates CRS seen in the clinic42,43,44. Although these models allow the study of specific immune cells in relation to CART efficacy, the humanized system is still primitive and needs further optimization. In addition, their use in studying off-target effects is limited since most of the host antigens on healthy tissue are murine in origin. Furthermore, the CD34+ humanized mouse model is technically challenging to develop45. However, human xenograft models in immunocompromised mice are commonly used to assess human CART cell efficacy prior to translating to clinical trials, and as demonstrated in this paper, they can be used to evaluate the toxicities associated with CART cell activity. However, the lack of immune system and immune cells such as T cells and NK cells limits their application for assessing off-tumor on-target toxicity, interactions of CART cells with innate immune cells, and CART cell inhibition by the tumor microenvironment.

The advantage of the protocol described here is that it requires simple procedures using NSG mice to observe and assess CART cell-associated toxicities. First, it is easy to reproduce, and it has been optimized to assess strategies to prevent CART toxicities through GM-CSF neutralization and to visualize CART cell expansion in correlation with CRS onset. When using this novel PDX model, it is important to correctly perform the critical steps, such as confirming the proper engraftment of the ALL blasts and the early monitoring of CRS onset right after CART administration. The assessment of cytokines and weight monitoring are crucial for determining CART-associated toxicities, and MRI defines the onset of NI. The downsides of this model are as follows: 1) as PDX ALL cells take longer to engraft and reach a high tumor burden, it requires approximately 2-3 months from tumor cell inoculation to CART19 cell treatment; 2) as a primary PDX model, there is significant patient to patient variability with regard to the time to engraftment and response to CART cell therapy; 3) tumor load cannot be assessed with bioluminescence imaging and requires frequent peripheral blood assessment; and 4) the lack of innate immune cells limits the ability of this model to study CART interactions with immune cells or the tumor microenvironment.

Interest in testing adoptive cells within more complex mouse models has grown over the past decade as it has become evident that simple immunodeficient mouse models are not ideal preclinical models. The PDX mouse model described here to assess CART cell-associated toxicities represents a promising preclinical model in the field of CART cell therapy as it mimics the timeline, symptoms, and markers of CRS and NI after CART cell infusion as seen in the clinic. Overall, the methodology described here provides a robust platform for assessing CART19 cell-associated toxicities as well as efficacy.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

S.S.K. is an inventor on patents in the field of CAR immunotherapy that are licensed to Novartis (through an agreement between Mayo Clinic, University of Pennsylvania, and Novartis) and to Mettaforge (through Mayo Clinic). R.L.S. and S.S.K. are inventors on patents in the field of CAR immunotherapy that are licensed to Humanigen. S.S.K. receives research funding from Kite, Gilead, Juno, Celgene, Novartis, Humanigen, MorphoSys, Tolero, Sunesis, Leahlabs, and Lentigen.

Acknowledgments

This work was partly supported through the National Institutes of Health (R37CA266344, 1K99CA273304), Department of Defense (CA201127), Mayo Clinic K2R pipeline (S.S.K.), the Mayo Clinic Center for Individualized Medicine (S.S.K.), and the Predolin Foundation (R.L.S.). In addition, we would like to thank the Mayo Clinic NMR Core Facility staff. Figure 1 was created in BioRender.com

Materials

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

DOWNLOAD MATERIALS LIST

References

  1. Turtle, C. J., et al. Immunotherapy of non-Hodgkin's lymphoma with a defined ratio of CD8+ and CD4+ CD19-specific chimeric antigen receptor-modified T cells. Science Translational Medicine. 8 (355), (2016).
  2. Kochenderfer, J. N., et al. Long-duration complete remissions of diffuse large B cell lymphoma after anti-CD19 chimeric antigen receptor T cell therapy. Molecular Therapy. 25 (10), 2245-2253 (2017).
  3. Kochenderfer, J. N., et al. Lymphoma remissions caused by anti-CD19 chimeric antigen receptor T cells are associated with high serum interleukin-15 levels. Journal of Clinical Oncology. 35 (16), 1803-1813 (2017).
  4. Maude, S. L., et al. Chimeric antigen receptor T cells for sustained remissions in leukemia. New England Journal of Medicine. 371 (16), 1507-1517 (2014).
  5. Park, J. H., et al. Long-term follow-up of CD19 CAR therapy in acute lymphoblastic leukemia. New England Journal of Medicine. 378 (5), 449-459 (2018).
  6. Kochenderfer, J. N., et al. Chemotherapy-refractory diffuse large B-cell lymphoma and indolent B-cell malignancies can be effectively treated with autologous T cells expressing an anti-CD19 chimeric antigen receptor. Journal of Clinical Oncology. 33 (6), 540-549 (2015).
  7. Anagnostou, T., Riaz, I. B., Hashmi, S. K., Murad, M. H., Kenderian, S. S. Anti-CD19 chimeric antigen receptor T-cell therapy in acute lymphocytic leukaemia: A systematic review and meta-analysis. Lancet Haematology. 7 (11), 816-826 (2020).
  8. Siegler, E. L., Kenderian, S. S. Neurotoxicity and cytokine release syndrome after chimeric antigen receptor T cell therapy: Insights into mechanisms and novel therapies. Frontiers in immunology. 11, 1973 (2020).
  9. Lee, D. W., et al. ASTCT consensus grading for cytokine release syndrome and neurologic toxicity associated with immune effector cells. Biology of Blood and Marrow Transplantation. 25 (4), 625-638 (2019).
  10. Maude, S. L., et al. Tisagenlecleucel in children and young adults with B-cell lymphoblastic leukemia. The New England Journal of Medicine. 378 (5), 439-448 (2018).
  11. Teachey, D. T., et al. Identification of predictive biomarkers for cytokine release syndrome after chimeric antigen receptor T-cell therapy for acute lymphoblastic leukemia. Cancer Discovery. 6 (6), 664-679 (2016).
  12. Neelapu, S. S., et al. Axicabtagene ciloleucel CAR T-cell therapy in refractory large B-cell lymphoma. The New England Journal of Medicine. 377 (26), 2531-2544 (2017).
  13. Locke, F. L., et al. Long-term safety and activity of axicabtagene ciloleucel in refractory large B-cell lymphoma (ZUMA-1): A single-arm, multicentre, phase 1-2 trial. The Lancet. Oncology. 20 (1), 31-42 (2019).
  14. Hunter, B. D., Jacobson, C. A. CAR T-cell associated neurotoxicity: Mechanisms, clinicopathologic correlates, and future directions. Journal of the National Cancer Institute. 111 (7), 646-654 (2019).
  15. Neelapu, S. S., et al. Axicabtagene ciloleucel CAR T-cell therapy in refractory large B-cell lymphoma. New England Journal of Medicine. 377 (26), 2531-2544 (2017).
  16. Schuster, S. J., et al. Tisagenlecleucel in adult relapsed or refractory diffuse large B-cell lymphoma. New England Journal of Medicine. 380 (1), 45-56 (2019).
  17. Santomasso, B. D., et al. Clinical and biological correlates of neurotoxicity associated with CAR T-cell therapy in patients with B-cell acute lymphoblastic leukemia. Cancer Discovery. 8 (8), 958-971 (2018).
  18. Lee, D. W., et al. Current concepts in the diagnosis and management of cytokine release syndrome. Blood. 124 (2), 188-195 (2014).
  19. Chen, F., et al. Measuring IL-6 and sIL-6R in serum from patients treated with tocilizumab and/or siltuximab following CAR T cell therapy. Journal of Immunological Methods. 434, 1-8 (2016).
  20. Ruff, M. W., Siegler, E. L., Kenderian, S. S. A concise review of neurologic complications associated with chimeric antigen receptor T-cell immunotherapy. Neurologic Clinics. 38 (4), 953-963 (2020).
  21. Sterner, R. M., Kenderian, S. S. Myeloid cell and cytokine interactions with chimeric antigen receptor-T-cell therapy: Implication for future therapies. Current Opinion in Hematology. 27 (1), 41-48 (2020).
  22. Gofshteyn, J. S., et al. Neurotoxicity after CTL019 in a pediatric and young adult cohort. Annals of Neurology. 84 (4), 537-546 (2018).
  23. Shalabi, H., et al. Systematic evaluation of neurotoxicity in children and young adults undergoing CD22 chimeric antigen receptor T-cell therapy. Journal of Immunotherapy. 41 (7), 350-358 (2018).
  24. Siegler, E. L., Wang, P. Preclinical models in chimeric antigen receptor-engineered T-cell therapy. Human Gene Therapy. 29 (5), 534-546 (2018).
  25. Mhaidly, R., Verhoeyen, E. Humanized mice are precious tools for preclinical evaluation of CAR T and CAR NK cell therapies. Cancers. 12 (7), 1915 (1915).
  26. Sakemura, R., et al. Development of a clinically relevant reporter for chimeric antigen receptor T-cell expansion, trafficking, and toxicity. Cancer Immunology Research. 9 (9), 1035-1046 (2021).
  27. Sterner, R. M., Cox, M. J., Sakemura, R., Kenderian, S. S. Using CRISPR/Cas9 to knock out GM-CSF in CAR-T cells. Journal of Visualized Experiments. (149), e59629 (2019).
  28. Sterner, R. M., et al. GM-CSF inhibition reduces cytokine release syndrome and neuroinflammation but enhances CAR-T cell function in xenografts. Blood. 133 (7), 697-709 (2019).
  29. Cox, M. J., et al. Leukemic extracellular vesicles induce chimeric antigen receptor T cell dysfunction in chronic lymphocytic leukemia. Molecular Therapy. 29 (4), 1529-1540 (2021).
  30. Sakemura, R., et al. Dynamic imaging of chimeric antigen receptor T cells with [18F]tetrafluoroborate positron emission tomography/computed tomography. Journal of Visualized Experiments. (180), e62334 (2022).
  31. Cox, M. J., Kenderian, S. S., et al. GM-CSF disruption in CART cells modulates T cell activation and enhances CART cell anti-tumor activity. Leukemia. 36 (6), 1635-1645 (2022).
  32. Pirko, I., Suidan, G. L., Rodriguez, M., Johnson, A. J. Acute hemorrhagic demyelination in a murine model of multiple sclerosis. Journal of Neuroinflammation. 5, 31 (2008).
  33. Denic, A., et al. MRI in rodent models of brain disorders. Neurotherapeutics. 8 (1), 3-18 (2011).
  34. Johnson, H. L., et al. CD8 T cell-initiated blood-brain barrier disruption is independent of neutrophil support. Journal of Immunology. 189 (4), 1937-1945 (2012).
  35. Johnson, H. L., et al. Perforin competent CD8 T cells are sufficient to cause immune-mediated blood-brain barrier disruption. PLoS One. 9 (10), 111401 (2014).
  36. Huggins, M. A., et al. Perforin expression by CD8 T cells is sufficient to cause fatal brain edema during experimental cerebral malaria. Infection and Immunity. 85 (5), 00985 (2017).
  37. Pennell, C. A., et al. Human CD19-targeted mouse T cells induce B cell aplasia and toxicity in human CD19 transgenic mice. Molecular Therapy. 26 (6), 1423-1434 (2018).
  38. Khadka, R. H., Sakemura, R., Kenderian, S. S., Johnson, A. J. Management of cytokine release syndrome: an update on emerging antigen-specific T cell engaging immunotherapies. Immunotherapy. 11 (10), 851-857 (2019).
  39. Sanmamed, M. F., Chester, C., Melero, I., Kohrt, H. Defining the optimal murine models to investigate immune checkpoint blockers and their combination with other immunotherapies. Annals of Oncology. 27 (7), 1190-1198 (2016).
  40. Mardiana, S., et al. A multifunctional role for adjuvant anti-4-1BB therapy in augmenting antitumor response by chimeric antigen receptor T cells. Cancer Research. 77 (6), 1296-1309 (2017).
  41. Pegram, H. J., et al. Tumor-targeted T cells modified to secrete IL-12 eradicate systemic tumors without need for prior conditioning. Blood. 119 (18), 4133-4141 (2012).
  42. Kalscheuer, H., et al. A model for personalized in vivo analysis of human immune responsiveness. Science Translational Medicine. 4 (125), (2012).
  43. Xia, J., et al. Modeling human leukemia immunotherapy in humanized mice. EBioMedicine. 10, 101-108 (2016).
  44. Holzapfel, B. M., Wagner, F., Thibaudeau, L., Levesque, J. P., Hutmacher, D. W. Concise review: humanized models of tumor immunology in the 21st century: Convergence of cancer research and tissue engineering. Stem Cells. 33 (6), 1696-1704 (2015).
  45. Cogels, M. M., et al. Humanized mice as a valuable pre-clinical model for cancer immunotherapy research. Frontiers in Oncology. 11, 784947 (2021).

Tags

Chimeric Antigen Receptor T Cell-Associated Toxicities Acute Lymphoblastic Leukemia Patient-Derived Xenograft Mouse Model CAR-T Toxicities CART19 Administration Clinic Observations Platform Translatable Model Clinically Relevant Model Human T-cells Tumor Cells Assessment Treatment Strategies Wellbeing Changes Motor Weakness Hunched Body Loss Of Body Weight Peripheral Blood Analysis Serum Isolation Cytokines MRI Imaging Gadolinium Saline Solution Insulin Syringe

Erratum

Formal Correction: Erratum: Assessment of Chimeric Antigen Receptor T Cell-Associated Toxicities Using an Acute Lymphoblastic Leukemia Patient-derived Xenograft Mouse Model
Posted by JoVE Editors on 03/14/2023. Citeable Link.

An erratum was issued for: Assessment of Chimeric Antigen Receptor T Cell-Associated Toxicities Using an Acute Lymphoblastic Leukemia Patient-derived Xenograft Mouse Model. The Authors section was updated from:

Claudia Manriquez Roman1,2,3,4,5
R. Leo Sakemura1,2
Fang Jin6
Roman H. Khadka6
Mohamad M. Adada1,2
Elizabeth L. Siegler1,2
Aaron J. Johnson6
Saad S. Kenderian1,2,6
1T Cell Engineering Laboratory, Mayo Clinic, Rochester,
2Division of Hematology, Mayo Clinic, Rochester,
3Mayo Clinic Graduate School of Biomedical Sciences, Mayo Clinic, Rochester,
4Department of Molecular Medicine, Mayo Clinic, Rochester,
5Regenerative Sciences PhD Program, Mayo Clinic, Rochester,
6Department of Immunology, Mayo Clinic, Rochester

to:

Claudia Manriquez Roman1,2,3,4,5
R. Leo Sakemura1,2
Brooke L. Kimball1,2
Fang Jin6
Roman H. Khadka6
Mohamad M. Adada1,2
Elizabeth L. Siegler1,2
Aaron J. Johnson6
Saad S. Kenderian1,2,6
1T Cell Engineering Laboratory, Mayo Clinic, Rochester,
2Division of Hematology, Mayo Clinic, Rochester,
3Mayo Clinic Graduate School of Biomedical Sciences, Mayo Clinic, Rochester,
4Department of Molecular Medicine, Mayo Clinic, Rochester,
5Regenerative Sciences PhD Program, Mayo Clinic, Rochester,
6Department of Immunology, Mayo Clinic, Rochester

Assessment of Chimeric Antigen Receptor T Cell-Associated Toxicities Using an Acute Lymphoblastic Leukemia Patient-Derived Xenograft Mouse Model
Play Video
PDF DOI DOWNLOAD MATERIALS LIST

Cite this Article

Manriquez Roman, C., Sakemura, R.… More

Manriquez Roman, C., Sakemura, R. L., Kimball, B. L., Jin, F., Khadka, R. H., Adada, M. M., Siegler, E. L., Johnson, A. J., Kenderian, S. S. Assessment of Chimeric Antigen Receptor T Cell-Associated Toxicities Using an Acute Lymphoblastic Leukemia Patient-Derived Xenograft Mouse Model. J. Vis. Exp. (192), e64535, doi:10.3791/64535 (2023).

Less
Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
Simple Hit Counter