JoVE Logo
Authors
Contact Us

Sign In

A subscription to JoVE is required to view this content. Sign in or start your free trial.

In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol outlines a detailed procedure for generating transient chimeric antigen receptor (CAR) T cells using non-integrating mRNA for cancer immunotherapy and provides reliable methods for evaluating CAR-T cells and their cytotoxic function.

Abstract

Chimeric antigen receptor (CAR) T cell therapy has emerged as a pioneering cancer treatment, achieving unprecedented success in treating certain hematological malignancies such as lymphomas and leukemias. However, as more cancer patients receive CAR-T cell therapies, treatment-associated secondary primary malignancies are increasingly being reported partly due to unexpected CAR transgene insertion, raising serious safety concerns. To address this issue, we describe here a nonviral, non-integrating approach to generate transient CAR-T cells using mRNA. We electroporated T cells with modified mRNA encoding a human epidermal growth factor receptor 2 (HER2)-specific CAR and generated transient HER2-targeted CAR-T cells. The CAR was efficiently expressed on the T cell surface 1 day after electroporation, increased by day 2, and dramatically declined by day 5. The transient CAR-T cells exhibited potent cytotoxicity against HER2-positive SKOV-3 ovarian cancer cells and secreted high levels of IFN-ϒ. This protocol provides a step-by-step guide for developing small-scale transient CAR-T cells without permanent CAR transgene integration, describing detailed procedures for preparation of CAR mRNA, activation and transfection of T cells, assessment of CAR expression, and in vitro analysis of CAR-T cell function. This method is suitable for transient CAR-T cell generation in both preclinical and clinical studies.

Introduction

Cancer is an increasingly important threat to human health, with an annually estimated 23.6 million newly diagnosed cases and 10 million deaths globally1. Surgical treatment combined with radiation and chemotherapy remains the gold standard of care for various types of localized noninvasive and invasive cancers2,3,4. Although the traditional systematic treatment has achieved tremendous success in managing early-stage cancers, it is very toxic and has limited effect on metastatic and hematological cancers5. Patients with these cancers undergo frequent treatments and endure significant toxicity in the hope of stabilizing and delaying disease progression. Chimeric antigen receptor (CAR) T cell therapy has recently emerged as a revolutionary targeted therapy that demonstrates potent efficacy for B cell hematological cancers and offers hope to these desperate patients6.

CAR-T cells are engineered T lymphocytes expressing a CAR on the cell surface that specifically recognizes tumor-associated antigens (TAAs) and activates the cells without requiring antigenic presentation from major histocompatibility complex proteins (MHCs)7. Upon activation, the CAR-T cells secrete a panel of cytokines and lyse tumor cells independently of MHCs, thus enhancing the destruction of tumors even when MHCs are downregulated or lost due to the immunosuppressive tumor microenvironment8,9. CARs consist of at least three distinct domains—an extracellular single-chain fragment variant (scFv) containing the antigen-recognizing variable regions of a TAA-specific antibody, a transmembrane domain that anchors the CAR to the cell surface, and an intracellular domain that mediates adaptor protein recruitment and downstream signaling10. Based on variations in the intracellular domains, the CAR structures have evolved through five generations so far, with the first generation containing only the CD3ζ activation domain and subsequent generations possessing one or more extra costimulatory domains from molecules such as 4-1BB and CD2811. These intracellular domains affect the CAR-T cell signaling mechanisms, proliferation, survival, and toxicity, which together determine the clinical antitumor efficacy. The clinical success of CAR-T cell therapy comes from second-generation CAR-T cells that specifically target CD19, a biomarker highly expressed on B-lineage cells, for treating B-cell leukemias and lymphomas12,13. To date, the Food and Drug Administration (FDA) has approved six second-generation CAR-T cell therapies targeting either CD-19 or the B-cell maturation antigen (BCMA) for relapsed or refractory hematological malignancies, including large B-cell lymphoma, mantle cell lymphoma, and multiple myeloma 6. The newer generations of CAR-T cells are currently undergoing intensive preclinical and clinical studies14,15.

Clinical production of CAR-T cells is a complex, expensive, and time-consuming process. Currently, the leukocytes used for CAR-T cell preparation come from the patients themselves to avoid allogeneic reactions, although donor-derived universal (or off-the-shelf) CAR-T cells are under active development and clinical evaluation16. Following isolation of the leukocytes from the patient's peripheral blood by leukapheresis, a foreign gene fragment encoding the CAR is introduced into the activated T cells using either viral or nonviral approaches17,18,19. Retroviruses and lentiviruses are the most common viral vectors for CAR gene delivery, which results in random and permanent integration of the CAR-encoding DNA into the T cell genome20. Nonviral approaches, including mRNA-lipid nanoparticles, transposon systems, and CRISPR/Cas9 genome editing, are less common21. Subsequently, the CAR-expressing T cells are expanded ex vivo on a large scale and then collected and reinfused back into the patients22,23. Manufacturing viral vectors that meet stringent Good Manufacturing Practice (GMP) standards are very difficult, complex, and costly, placing a significant burden on manufacturers, regulatory agencies, and patients, thereby limiting its widespread clinical application.

Despite the exciting success of CAR-T therapy, there is growing concern about its safety. Given the random nature of CAR transgene integration into the T cell genome during viral or transposon transduction, disruption of tumor suppressor genes or activation of oncogenes may occur, leading to malignant transformation of the T cells24,25. Since the FDA's approval of the first CAR-T cell therapy product in 2017, follow-up studies show that as many as 15% of patients who receive the therapy develop secondary primary malignancies (SPMs) such as T cell lymphoma (TCL), non-small cell lung cancer (NSCLC) and skin cancer26,27,28,29. Strikingly, CAR transgenes were highly detected in some SPMs30,31, and some cases of TCL were directly caused by abnormal expansion of the CAR-T cells24,32,33, indicating a direct role of the CAR-T products in inducing some SPMs. Other studies further identified CAR transgene insertions in critical genes such as TET234, JAK135, and PBX236 in the SPM tumor cells. In light of the accumulating evidence of T-cell malignancies following CAR-T cell therapy, the FDA recently concluded that a boxed warning highlighting a serious risk of T-cell malignancy is required for all currently approved CD19-directed and BCMA-targeted CAR-T cell therapies37. However, large-cohort studies following the long-term effects of CAR-T treatment report an incidence of SPMs ranging from 3.8% to 15% among treated patients26,27,28,38,39. The incidence is not significantly higher than that observed in blood cancer patients receiving traditional systematic treatment40,41, suggesting a relatively safe profile for CAR-T cell therapy. Nonetheless, there is an urgent need to develop safe and efficacious CAR-T cells that minimize the risk of SPMs through more cost-effective approaches.

Here, we describe a detailed protocol for a nonviral, non-integrating approach to generate CAR-T cells. The goal is to provide a straightforward, step-by-step guide for generating small-scale, effective CAR-T cells with transient CAR expression, thereby avoiding insertional mutagenesis and minimizing the risk of SPMs. We show that the electroporation of T cells with modified mRNA encoding a CAR specific to the TAA HER2 resulted in robust and transient expression of the anti-HER2 CAR on the T cell surface. While expressing the CAR, the T cells potently killed HER2-positive tumor cells and secreted a high level of IFN-ϒ. The protocol is described in four separate but continuous sections that include preparation of CAR mRNA, activation and electroporation of T cell, assessment of CAR expression, and analysis of CAR-T cell function. This approach is suitable for developing and producing transient CAR-T cells for both academic research and clinical cell therapies.

Protocol

The PBMCs used here were previously isolated from whole blood from the Stanford Hospital Blood Center according to the Institutional Review Board (IRB)-approved protocol (13942) using the Ficoll-Paque density gradient described before42. The participants' informed consent was not applicable as the blood we used to collect the PBMCs was obtained commercially from the Stanford Hospital Blood Center.

1. Preparation of CAR mRNA

  1. Template preparation
    NOTE: Either a linearized plasmid or a PCR fragment can serve as a template to synthesize CAR mRNA in vitro, provided they contain a proper 5' SP6 or T7 promoter, a 5' untranslated region (UTR), a CAR transgene, and a 3'UTR with an appropriate poly(A) tail. In this study, we used a CAR plasmid DNA construct containing an SP6 promoter, a short human beta-globin 5'UTR, a CAR transgene, and a 3'UTR composed of two tandem beta-globin 3'UTRs followed by a 112-bp poly(A) tail43. The UTRs, coding sequence, and poly(A) tail all affect RNA stability and translational efficiency43,44,45. A poly(A) tail of 120 bp has been shown to increase protein yield in cell culture compared to shorter poly(A) tails43.
    1. Prepare CAR construct plasmid DNA using a plasmid prep kit according to the manufacturer's instructions. High-quality plasmid DNA is essential for generating a high yield of CAR mRNA.
    2. Linearize the CAR plasmid DNA with the restriction enzyme BglII. Choose a restriction enzyme that cuts after the poly(A) tail, preferentially close to the end of the poly(A) tail. Digest 10 µg (or 10,000 ng) of the plasmid quantified by a spectrophotometer in a 100 µL reaction mix containing 10 U of the enzyme, 10 µL of digestion buffer, and nuclease-free water overnight at 37 °C.
      NOTE: Make sure the restriction enzyme does not cut within the CAR transgene and its flanking promotor, UTR, and poly(A) tail sequences. Complete linearization of the CAR plasmid is necessary for efficient CAR mRNA transcription.
    3. Add 200 µg/mL protease K and 0.5% w/v SDS into the digestion mix and incubate at 50 °C for 1 h.
    4. Purify the linearized CAR plasmid DNA. Add an equal volume of phenol: chloroform: isoamyl alcohol (25:24:1, v/v) to the digestion reaction. Vortex vigorously for approximately 15 s. Spin the sample in a bench-top centrifuge at maximum speed for 5 min at room temperature (RT).
    5. Transfer the upper aqueous layer, which contains the DNA, to a new microcentrifuge tube. Avoid transfer of any of the middle and bottom layers, which contain protein and phenol impurities. Perform a second round of extraction with chloroform to reduce impurities.
    6. Add 1/10th volume of 7.5 M ammonium acetate or 3 M sodium acetate and mix gently. Add 2.5 volumes of ethanol to precipitate the DNA. Store the sample at −20 °C for at least 30 min.
    7. Spin the sample at maximum speed for 15 min at 4 °C to pellet the DNA. Remove the supernatant carefully. Add 500 µL of 70% ethanol to wash the DNA pellet.
    8. Spin the sample again for 2 min at 4 °C. Carefully remove the supernatant. Leave a small amount of solution in the tube.
    9. Repeat the spin for 2 min at 4°C. Carefully remove the remaining supernatant. Air-dry the DNA pellet at RT for about 10 min in a sterile hood until no ethanol solution is visible at the bottom of the tube.
      NOTE: Avoid overdrying the DNA pellet, as it might be difficult to dissolve.
    10. Add 20 µL of nuclease-free water and pipet up and down gently to dissolve the DNA. Centrifuge the sample briefly at maximum speed. Measure DNA concentration with a spectrophotometer.
    11. Store the sample on ice for immediate use or at −20 °C for future use. Load 100 ng of the purified DNA on an agarose gel and verify the size and integrity. The DNA should appear as a single, distinct band of the expected size. No extra or smeared bands should be observed.
      ​NOTE: The quality of the template DNA is critical for efficient mRNA transcription. Avoid using impure, fragmented, and degraded DNA as a template for mRNA transcription.
  2. In vitro transcription (IVT)
    1. Synthesize the CAR mRNA in vitro. Make sure to use nuclease-free reagents, tubes, and pipette tips throughout the procedures.
    2. Thaw the reagents listed in Table 1 to RT, except for the SP6 RNA polymerase mix, which should be stored at −20 °C till use. Prepare the transcription reaction in the order shown in Table 1 at RT. Mix gently and incubate at 37 °C for at least 2 h.
      NOTE: The reaction can be scaled up or down based on the amount of mRNA needed. The reaction shown above typically yields at least 100 µg (or 100,000 ng) of capped mRNA when using high-quality template DNA. Longer incubation, i.e., overnight, may increase mRNA yield.
    3. Add 2 µL of RNase-free DNase I to the reaction, mix gently, and incubate for 15 min at 37 °C to degrade the template DNA.
    4. Purify the transcribed CAR mRNA using an RNA Cleanup Kit (see Table of Materials). Follow the kit manufacturer's instructions for the purification. Elute the mRNA in nuclease-free water.
      NOTE: The mRNA can also be purified using the phenol-chloroform extraction method shown above (step 1.1.4).
    5. Measure the mRNA concentration with a spectrophotometer. Load at least 200 ng of the mRNA on an agarose gel and verify its size and integrity. The mRNA should appear as a dominant band of the expected size, without any additional or smeared bands. Store the mRNA at −80 °C until use.

2. Activation and electroporation of T Cell

  1. Suspend 1 x 106 cryopreserved human peripheral blood mononuclear cells (PBMCs) in 1 mL of CAR-T medium (AIM-V medium supplemented with 10% fetal bovine serum (FBS), penicillin/streptomycin, and 10 ng/ml of human recombinant IL-2). Activate the T cells with an equal number of prewashed Human T-Activator CD3/CD28 beads (see Table of Materials). Culture and expand the cells at 37 °C in a humidified incubator with 5% CO2 for several days.
    NOTE: The amount of PBMCs can be scaled up or down based on the amount of T cells required for subsequent experiments.
  2. Thaw prepared CAR mRNA on ice. Add 1 mL of CAR-T medium per well to two wells of a 12-well plate and equilibrate the medium by placing the plate in the incubator at 37 °C.
  3. Transfer 5 x 106 T cells to a sterile tube and remove the beads by putting the tube onto a magnetic stand and carefully pipetting the cell suspension to a new tube.
  4. Centrifuge the tube at 300 x g for 5 min at RT. Discard the supernatant and resuspend the cell pellet in 1 mL of sterile PBS.
  5. Centrifuge the tube again. Discard the supernatant and resuspend the cells in 200 µL of Neon buffer T (see Table of Materials).
  6. Pipet 100 µL of cells to a new tube and add 5 µg (or 5,000 ng) of CAR mRNA to the cells46. Supplement Neon buffer T to a total of 125 µL and mix by pipetting up and down.
    NOTE: The amount of CAR mRNA to be transfected by electroporation may be increased up to 8 µg (or 8,000 ng) per 1 x 106 T cells without negatively affecting cell viability, and a higher amount of mRNA may lead to increased CAR expression46.
  7. Add 25 µL of Neon buffer T to the remaining 100 µL cells. This serves as the negative control.
  8. Electroporate the T cells. Use the Neon Pipette to aspirate the T cells/CAR mRNA mixture or the negative control T cells into a Neon 100 µL tip and electroporate the cells with a 10 ms pulse of 1800 V.
  9. Immediately transfer the cells to the equilibrated culture medium in the 12-well plate (prepared at step 2.2) after the electroporation.
  10. Return the plate back to the incubator and expand the cells until ready for use. To ensure healthy and rapid growth of the CAR T cells, maintain cell density between 1−3 x 106 cells/mL with fresh medium until needed.

3. Assessment of CAR expression

  1. Mix the CAR T cells by pipetting up and down gently several times in each well, and transfer at least 1 x 105 cells to a 5 mL FACS tube.
  2. Add 3 mL of cold wash buffer (PBS containing 2 mM EDTA and 0.5% w/v BSA) to the tube. Centrifuge the tube at 500 x g for 5 min at RT. Remove the supernatant carefully.
  3. Resuspend the cell pellet in 100 µL of wash buffer. Add 2 µL of the CAR detection reagent allophycocyanin (APC)-conjugated goat anti-human IgG F(ab')2 antibody (see Table of Materials) and 2 µL of a viability dye 7-AAD into the suspended cells.
    NOTE: The CAR can also be detected using an appropriately labeled target antigen or an anti-tag antibody if the CAR contains a tag (i.e., FLAG). If an unlabeled antibody is used for CAR detection, incubation with an appropriate fluorescence-labeled secondary antibody is required after step 3.4.
  4. Mix the sample by gently shaking the tube and incubate on ice for 30 min. Avoid light during the incubation. Wash the cells with 3 mL of cold wash buffer, and centrifuge the tube at 500 x g for 5 min at RT.
  5. Discard the supernatant and resuspend the cells in 100 µL of wash buffer. Analyze the cells using a flow cytometer immediately. Make sure the detection reagent's fluorescent label and the viability dye are properly compensated using appropriate positive and negative controls.
  6. Evaluate the CAR expression using the following gating strategy: Gate on the cells using the FSC-A vs SSC-A plot; Gate on the single cells using the FSC-A vs FSC-H plot; Gate on the live cells using the 7-AAD staining.
  7. Calculate the percentage of live T cells that were stained by the anti-human IgG F(ab')2 antibody. Compare the CAR-T cells to the negative control T cells, which should have minimal staining.

4. Analysis of CAR-T cell function

  1. Cytotoxicity analysis
    1. Make sure the target tumor cells express the antigen in a conformation that can be recognized by the CAR. To confirm the antigen expression, analyze the tumor cells by flow cytometry using the parent antibody from which the CAR was derived. If the CAR cannot recognize the tumor cells, then the CAR-T cells will fail to kill the tumor cells.
    2. Prepare target tumor cells expressing the CAR's antigen on the cell surface. Trypsinize the adherent tumor cells with 0.05% trypsin-EDTA solution for 1 min at 37 °C and prepare single cell suspension with appropriate culture medium at 1−4 x 105 cells/mL.
      NOTE: The tumor cells should be well-adherent to ensure accurate analysis in the subsequent steps. If the tumor cells adhere poorly, detach easily or grow in suspension, coat the wells of the plate used in the following step with an appropriate antibody, polycationic polymer, or biological material such as fibronectin or collagen to facilitate cell attachment.
    3. Prepare an E-Plate (see Table of Materials). Design the experimental setup. Add 50 µL of pre-warmed tumor cell culture medium into each well and place the E-Plate into the Real-Time Cell Analysis (RTCA) instrument to measure background impedance. Measure the impedance at 1 min per measurement (or sweep) for 3 sweeps.
      NOTE: The background impedance is used to set the cell index to zero. Without background impedance measurement, the wells cannot be compared to each other, rendering the results non-interpretable.
    4. Remove the E-Plate from the RTCA instrument and add 100 µL of the tumor cell suspension to each well. Therefore, each well contains 1−4 x 104 cells in 150 µL total volume.
      NOTE: The number of tumor cells to be seeded in each well should be determined prior to the experiment. Seed the E-Plate with a range of cell numbers, then monitor the cell indexes over the next 24−48 hours. The desired number of cells maintains a steady increase in cell index for at least 24 hours without plateauing. Excessive cell seeding can accelerate nutrient depletion and medium acidification, resulting in early plateauing of the cell index. In addition, replicate wells should be included for each group to identify potential outlier measurements.
    5. Equilibrate the E-Plate at RT for 30 min. This step allows the tumor cells to settle and adhere to the bottom of the wells.
      NOTE: This step is necessary for accurate cell index measurement. Skipping this step usually causes well-to-well variability in cell index measurements and uneven tumor cell adhesion within the wells, resulting in increased noise in the assay.
    6. Place the E-Plate back to the RTCA instrument and monitor the cell indexes for about 24 h, with sweeps taken every 5−15 min.
    7. The next day, prepare the CAR-T cell suspension. Suspend the CAR-T cells (the effector cells) from step 2.10 in RPMI 1640 medium plus 10% FBS to a concentration determined by the tumor cell (the target cell) and the desired effector:target (E:T) cell ratio. For example, if the tumor cell suspension was at a density of 2 x 105 cells/mL and an E:T ratio of 10:1 is desired, then the CAR-T cells should be suspended to 2 x 106 cells/mL. Include proper controls such as tumor cells alone (no effector cells) and control T cells (T cells electroporated without mRNA or with control mRNA).
      NOTE: For optimal results, use the CAR-T cells 1-2 days after electroporation when the surface CAR expression is high. To explore the optimal E:T ratio, a range of E:T ratios (for example, 3:1 to 20:1) can be tested. Unless the CAR-T frequency is very low, i.e., under 10%, an E:T ratio of 10:1 or 20:1 is usually sufficient to achieve near-complete killing of the tumor cells.
    8. Pause the cell index recording and remove the E-Plate from the RTCA instrument.
    9. Tilt the plate slightly, remove 50 µL of culture medium from each well carefully without touching the tumor cells, and add 100 µL of the CAR-T cells or control T cells into each well.
    10. Return the E-Plate to the RTCA instrument and resume cell index monitoring for at least 24 h with sweeps taken every 5−15 min.
      NOTE: The length of cell index monitoring depends on how quickly the CAR-T cells target the tumor cells. A 24 h incubation period is usually sufficient for the CAR-T cells to exert maximum cytotoxic effect.
    11. Stop the cell index monitoring, remove the E-Plate from the RTCA instrument, and analyze the cytotoxicity result. Calculate cytotoxicity as the percentage of tumor cells that were killed by the CAR-T cells. Calculate mean cytotoxicity of the CAR-T cells and control T cells using the RTCA software. To calculate the cytotoxicity manually, use the formula
      ​cytotoxicity = ((cell index of tumor cells) − (cell index of tumor cells plus effector cells)) / (cell index of tumor cells) × 100%.
  2. Cytokine secretion analysis
    1. To analyze the cytokines such as IFN-ϒ secreted by the CAR-T cells during the cytotoxicity analysis, transfer the supernatant from the E-Plate to a round-bottom or V-shaped 96-well plate.
      NOTE: If a time-course analysis of the cytokine secretion is desired, pause the cell index monitoring at step 4.1.10, collect a small portion of the supernatant from each well, return the E-Plate back to the RTCA instrument, and resume the monitoring.
    2. Centrifuge the 96-well plate at 300 x g for 5 min at RT. Carefully transfer the supernatants to a new 96-well plate without disturbing the cell pellets.
    3. Measure the cytokine levels in the supernatants using commercial enzyme-linked immunosorbent assay (ELISA) kits according to the manufacturer's instructions. The supernatants usually require dilution before measurement, sometimes over 100-fold, depending on the cytokines being analyzed.
    4. Analyze the data by interpolating the sample absorbance values against a standard curve generated from serially diluted standards. Prepare graphs with error bars representing the mean ± standard deviation (SD) using appropriate software. To compare the difference between treatments, a Student's t test can be performed. A p-value less than 0.05 is usually considered statistically significant.
      NOTE: Data points that are clear outliers, likely due to experimental errors, should be excluded from the analysis.

Results

We constructed a second-generation HER2-targeted CAR containing a scFv derived from the humanized anti-HER2 mouse monoclonal antibody (mAb) 4D547, a transmembrane region, and an intracellular 4-1BB costimulatory domain followed by the CD3ζ activation domain (Figure 1A). The DNA sequence encoding the CAR contained a 5' SP6 promoter to drive transcription of the CAR mRNA in vitro (Figure 1B). The CAR mRNA was approximately...

Discussion

In this study, we describe a detailed nonviral, non-integrating approach for generating transient CAR-T cells and provide technical procedures for assessing CAR-T cell function. This approach avoids the use of conventional retroviral and lentiviral vectors for permanent and random CAR transgene delivery, instead leveraging electroporation to efficiently introduce in vitro modified CAR mRNA into T cells. This method enables high-level, transient CAR expression on the T cell surface, significantly reducing the ris...

Disclosures

The authors Liang Hu, Robert Berahovich, Yanwei Huang, Shiming Zhang, Jinying Sun, Xianghong Liu, Hua Zhou, Shirley, Xu, Haoqi Li, and Vita Golubovskaya are employees of ProMab Biotechnologies. Lijun Wu is an employee and shareholder of ProMab Biotechnologies.

Acknowledgements

This work was supported by ProMab Biotechnologies.

Materials

NameCompanyCatalog NumberComments
7-AAD viability dye Biolegend420404
ACEA Novocyte flow cytometerAgilentNovoCyte 3000
AIM-V mediumGibco12055083
APC goat anti-human IgG F(ab')2 antibodyJackson ImmunoResearch Laboratories109-136-097
BglII Restriction EnzymeNew England BioLabs (NEB)R0144L
3´-O-Me-m7G(5')ppp(5')G RNA Cap Structure AnalogNEBS1411S
DMEM, high glucoseGibco11965092
Dynabeads Human T-Activator CD3/CD28 beadsGibco11131D
E-Plate 96Agilent5232376001
EthanolSigma459844-4L
FBSLonza.com14-503F
HiScribe SP6 RNA Synthesis KitNew England BioLabs (NEB)E2070S
Human IL-2 Recombinant ProteinGibco15140122
Millennium RNA MarkersInvitrogenAM7150
Monarch RNA Cleanup Kit (500 μg)NEBT2050L
N1-Methylpseudo-UTPTrilinkN-1081-10
Neon Transfection InstrumentInvitrogenMPK5000
Neon Transfection System 100-μL KitInvitrogenMPK10096
Penicillin-Streptomycin (10000 U/mL)Gibco14-503F
RPMI 1640 MediumGibco11875135
Trypsin-EDTA (0.05%), phenol redGibco25300120
UltraPure Phenol:Chloroform:Isoamyl Alcohol (25:24:1, v/v)Invitrogen15593049
xCELLigence Real-Time Cell Analysis (RTCA) instrumentAgilentRTCA MP
ZymoPURE II Plasmid Midiprep KitZymo ResearchD4201

References

  1. Ataeinia, B., et al. National and subnational incidence, mortality, and years of life lost due to breast cancer in iran: Trends and age-period-cohort analysis since 1990. Front Oncol. 11, 561376 (2021).
  2. Jordan, R. M., Oxenberg, J. . Breast cancer conservation therapy. , (2024).
  3. Tannock, I. F. Combined modality treatment with radiotherapy and chemotherapy. Radiother Oncol. 16 (2), 83-101 (1989).
  4. Mcree, A. J., Cowherd, S., Wang, A. Z., Goldberg, R. M. Chemoradiation therapy in the management of gastrointestinal malignancies. Future Oncol. 7 (3), 409-426 (2011).
  5. Palumbo, M. O., et al. Systemic cancer therapy: Achievements and challenges that lie ahead. Front Pharmacol. 4, 57 (2013).
  6. Mitra, A., et al. From bench to bedside: The history and progress of car t cell therapy. Front Immunol. 14, 1188049 (2023).
  7. Lim, W. A., June, C. H. The principles of engineering immune cells to treat cancer. Cell. 168 (4), 724-740 (2017).
  8. Silveira, C. R. F., et al. Cytokines as an important player in the context of car-t cell therapy for cancer: Their role in tumor immunomodulation, manufacture, and clinical implications. Front Immunol. 13, 947648 (2022).
  9. Labanieh, L., Majzner, R. G., Mackall, C. L. Programming car-t cells to kill cancer. Nat Biomed Eng. 2 (6), 377-391 (2018).
  10. Sadelain, M., Brentjens, R., Riviere, I. The basic principles of chimeric antigen receptor design. Cancer Discov. 3 (4), 388-398 (2013).
  11. Tokarew, N., Ogonek, J., Endres, S., Von Bergwelt-Baildon, M., Kobold, S. Teaching an old dog new tricks: Next-generation car t cells. Br J Cancer. 120 (1), 26-37 (2019).
  12. Kochenderfer, J. N., et al. Eradication of b-lineage cells and regression of lymphoma in a patient treated with autologous t cells genetically engineered to recognize cd19. Blood. 116 (20), 4099-4102 (2010).
  13. Brentjens, R. J., et al. Safety and persistence of adoptively transferred autologous cd19-targeted t cells in patients with relapsed or chemotherapy refractory b-cell leukemias. Blood. 118 (18), 4817-4828 (2011).
  14. Tomasik, J., Jasinski, M., Basak, G. W. Next generations of car-t cells - new therapeutic opportunities in hematology. Front Immunol. 13, 1034707 (2022).
  15. Asmamaw Dejenie, T., et al. Current updates on generations, approvals, and clinical trials of car t-cell therapy. Hum Vaccin Immunother. 18 (6), 2114254 (2022).
  16. Sun, W., Jiang, Z., Jiang, W., Yang, R. Universal chimeric antigen receptor t cell therapy - the future of cell therapy: A review providing clinical evidence. Cancer Treat Res Commun. 33, 100638 (2022).
  17. Wang, X., Riviere, I. Clinical manufacturing of car t cells: Foundation of a promising therapy. Mol Ther Oncolytics. 3, 16015 (2016).
  18. Pinto, I. S., Cordeiro, R. A., Faneca, H. Polymer- and lipid-based gene delivery technology for car t cell therapy. J Control Release. 353, 196-215 (2023).
  19. Magnani, C. F., et al. Sleeping beauty-engineered car t cells achieve antileukemic activity without severe toxicities. J Clin Invest. 130 (11), 6021-6033 (2020).
  20. Irving, M., Lanitis, E., Migliorini, D., Ivics, Z., Guedan, S. Choosing the right tool for genetic engineering: Clinical lessons from chimeric antigen receptor-t cells. Hum Gene Ther. 32 (19-20), 1044-1058 (2021).
  21. Moretti, A., et al. The past, present, and future of non-viral car t cells. Front Immunol. 13, 867013 (2022).
  22. Song, H. W., et al. Car-t cell expansion platforms yield distinct t cell differentiation states. Cytotherapy. 26 (7), 757-768 (2024).
  23. Stephenson, M., Grayson, W. Recent advances in bioreactors for cell-based therapies. F1000Res. 7, (2018).
  24. Micklethwaite, K. P., et al. Investigation of product-derived lymphoma following infusion of piggybac-modified cd19 chimeric antigen receptor t cells. Blood. 138 (16), 1391-1405 (2021).
  25. Ghilardi, G., et al. T cell lymphoma and secondary primary malignancy risk after commercial car t cell therapy. Nat Med. 30 (4), 984-989 (2024).
  26. Steffin, D. H. M., et al. Long-term follow-up for the development of subsequent malignancies in patients treated with genetically modified iecs. Blood. 140 (1), 16-24 (2022).
  27. Cappell, K. M., et al. Long-term follow-up of anti-cd19 chimeric antigen receptor t-cell therapy. J Clin Oncol. 38 (32), 3805-3815 (2020).
  28. Cordeiro, A., et al. Late events after treatment with cd19-targeted chimeric antigen receptor modified t cells. Biol Blood Marrow Transplant. 26 (1), 26-33 (2020).
  29. Zhao, A., et al. Secondary myeloid neoplasms after cd19 car t therapy in patients with refractory/relapsed b-cell lymphoma: Case series and review of literature. Front Immunol. 13, 1063986 (2022).
  30. Verdun, N., Marks, P. Secondary cancers after chimeric antigen receptor t-cell therapy. N Engl J Med. 390 (7), 584-586 (2024).
  31. Ruella, M., et al. Induction of resistance to chimeric antigen receptor t cell therapy by transduction of a single leukemic b cell. Nat Med. 24 (10), 1499-1503 (2018).
  32. Bishop, D. C., et al. Development of car t-cell lymphoma in 2 of 10 patients effectively treated with piggybac-modified cd19 car t cells. Blood. 138 (16), 1504-1509 (2021).
  33. Shah, N. N., et al. Clonal expansion of car t cells harboring lentivector integration in the cbl gene following anti-cd22 car t-cell therapy. Blood Adv. 3 (15), 2317-2322 (2019).
  34. Fraietta, J. A., et al. Disruption of tet2 promotes the therapeutic efficacy of cd19-targeted t cells. Nature. 558 (7709), 307-312 (2018).
  35. Heinrich, T., et al. Mature t-cell lymphomagenesis induced by retroviral insertional activation of janus kinase 1. Mol Ther. 21 (6), 1160-1168 (2013).
  36. Harrison, S. J., et al. Car+ t-cell lymphoma post ciltacabtagene autoleucel therapy for relapsed refractory multiple myeloma. Blood. 142 (Supplement 1), 6939-6939 (2023).
  37. FDA Administration. . Fda requires boxed warning for t cell malignancies following treatment with bcma-directed or cd19-directed autologous chimeric antigen receptor (car) t cell immunotherapies. , (2024).
  38. Elsallab, M., et al. Second primary malignancies after commercial car t-cell therapy: Analysis of the fda adverse events reporting system. Blood. 143 (20), 2099-2105 (2024).
  39. Levine, B. L., et al. Unanswered questions following reports of secondary malignancies after car-t cell therapy. Nat Med. 30 (2), 338-341 (2024).
  40. Tward, J. D., Wendland, M. M., Shrieve, D. C., Szabo, A., Gaffney, D. K. The risk of secondary malignancies over 30 years after the treatment of non-hodgkin lymphoma. Cancer. 107 (1), 108-115 (2006).
  41. Kumar, V., et al. Trends in the risk of second primary malignancies among survivors of chronic lymphocytic leukemia. Blood Cancer J. 9 (10), 75 (2019).
  42. Holtkamp, S., et al. Modification of antigen-encoding rna increases stability, translational efficacy, and t-cell stimulatory capacity of dendritic cells. Blood. 108 (13), 4009-4017 (2006).
  43. Dvir, S., et al. Deciphering the rules by which 5'-utr sequences affect protein expression in yeast. Proc Natl Acad Sci U S A. 110 (30), E2792-E2801 (2013).
  44. Van Der Velden, A. W., Voorma, H. O., Thomas, A. A. Vector design for optimal protein expression. Biotechniques. 31 (3), 572-580 (2001).
  45. Jaatinen, T., Laine, J. Isolation of mononuclear cells from human cord blood by ficoll-paque density gradient. Curr Protoc Stem Cell Biol. Chapter 2 (Unit 2A), 1 (2007).
  46. Zhao, Y., et al. High-efficiency transfection of primary human and mouse t lymphocytes using rna electroporation. Mol Ther. 13 (1), 151-159 (2006).
  47. Carter, P., et al. Humanization of an anti-p185her2 antibody for human cancer therapy. Proc Natl Acad Sci U S A. 89 (10), 4285-4289 (1992).
  48. Hu, L., et al. Her2-cd3-fc bispecific antibody-encoding mrna delivered by lipid nanoparticles suppresses her2-positive tumor growth. Vaccines. 12 (7), 808 (2024).
  49. Perales-Puchalt, A., et al. DNA-encoded bispecific t cell engagers and antibodies present long-term antitumor activity. JCI Insight. 4 (8), e126086 (2019).
  50. Schwanhausser, B., et al. Global quantification of mammalian gene expression control. Nature. 473 (7347), 337-342 (2011).
  51. Kozak, M. Features in the 5' non-coding sequences of rabbit alpha and beta-globin mrnas that affect translational efficiency. J Mol Biol. 235 (1), 95-110 (1994).
  52. Wu, Q., et al. Translation affects mrna stability in a codon-dependent manner in human cells. Elife. 8, e45396 (2019).
  53. Henderson, J. M., et al. Cap 1 messenger rna synthesis with co-transcriptional cleancap((r)) analog by in vitro transcription. Curr Protoc. 1 (2), e39 (2021).
  54. Kariko, K., et al. Incorporation of pseudouridine into mrna yields superior nonimmunogenic vector with increased translational capacity and biological stability. Mol Ther. 16 (11), 1833-1840 (2008).
  55. Gehl, J. Electroporation: Theory and methods, perspectives for drug delivery, gene therapy and research. Acta Physiol Scand. 177 (4), 437-447 (2003).
  56. Stacey, M., Fox, P., Buescher, S., Kolb, J. Nanosecond pulsed electric field induced cytoskeleton, nuclear membrane and telomere damage adversely impact cell survival. Bioelectrochemistry. 82 (2), 131-134 (2011).
  57. Meaking, W. S., Edgerton, J., Wharton, C. W., Meldrum, R. A. Electroporation-induced damage in mammalian cell DNA. Biochim Biophys Acta. 1264 (3), 357-362 (1995).
  58. Zhang, J., et al. Non-viral, specifically targeted car-t cells achieve high safety and efficacy in b-nhl. Nature. 609 (7926), 369-374 (2022).
  59. Von Auw, N., et al. Comparison of two lab-scale protocols for enhanced mrna-based car-t cell generation and functionality. Sci Rep. 13 (1), 18160 (2023).
  60. Gauthier, J., Turtle, C. J. Insights into cytokine release syndrome and neurotoxicity after cd19-specific car-t cell therapy. Curr Res Transl Med. 66 (2), 50-52 (2018).
  61. Beatty, G. L., et al. Activity of mesothelin-specific chimeric antigen receptor t cells against pancreatic carcinoma metastases in a phase 1 trial. Gastroenterology. 155 (1), 29-32 (2018).
  62. Beatty, G. L., et al. Mesothelin-specific chimeric antigen receptor mrna-engineered t cells induce anti-tumor activity in solid malignancies. Cancer Immunol Res. 2 (2), 112-120 (2014).
  63. Tchou, J., et al. Safety and efficacy of intratumoral injections of chimeric antigen receptor (car) t cells in metastatic breast cancer. Cancer Immunol Res. 5 (12), 1152-1161 (2017).
  64. Wu, J., Wu, W., Zhou, B., Li, B. Chimeric antigen receptor therapy meets mrna technology. Trends Biotechnol. 42 (2), 228-240 (2024).

Reprints and Permissions

Request permission to reuse the text or figures of this JoVE article

Request Permission

Explore More Articles

Nonviral ApproachChimeric Antigen Receptor T CellsCAR T Cell TherapyMRNACancer ImmunotherapyTransient CAR T CellsElectroporationGene IntegrationViral TransductionSafety ConcernsManufacturing ProceduresHuman Peripheral Blood Mononuclear CellsCD3 CD28 BeadsIn Vitro TranscriptionRNA Cleanup Kit

This article has been published

Video Coming Soon

JoVE Logo

Privacy

Terms of Use

Policies

Contact Us

Recommend to library

JoVE NEWSLETTERS

Research

Education

Authors

Librarians

ABOUT JoVE

Copyright © 2025 MyJoVE Corporation. All rights reserved