shRNA-based PD-1 suppression preserves memory phenotype and function of CD19-targeted CAR-T cell

Introduction

Chimeric antigen receptor T cell (CAR-T) therapy is a promising approach in cancer immunotherapy, wherein patient-derived T cells are engineered to express CARs targeting tumor-associated antigens (1). A typical CAR consists of an extracellular antigen-binding domain, hinge, transmembrane domain, and intracellular signaling domain (e.g., CD3ζ and co-stimulatory domain) (2,3). Upon antigen recognition, CAR signaling triggers T cell proliferation and effector responses, enabling tumor cell lysis (1).

Cluster of differentiation 19 (CD19)-targeted CAR-T cells have shown impressive efficacy in treating B-cell acute lymphoblastic leukemia (B-ALL), with initial complete remission rates reaching ~81% (4,5). However, long-term responses remain suboptimal, with durable remissions observed in only ~50% of patients (4,6,7). A major contributor to this limitation is CAR-T cell exhaustion, a dysfunctional state characterized by impaired proliferative capacity, reduced cytotoxicity, and diminished persistence (8). Within the immunosuppressive tumor microenvironment, sustained antigen stimulation and chronic activation drive memory-like CAR-T cells toward an exhausted phenotype, ultimately compromising their therapeutic potency (9).

The programmed cell death protein 1 (PD-1)/programmed death ligand 1 (PD-L1) immune checkpoint axis plays a key role in mediating T cell exhaustion (10). PD-1, which is upregulated upon chronic activation, transduces inhibitory signals upon binding to its ligand PD-L1. This interaction impairs effector function by suppressing multiple downstream signaling pathways, including the Lck-ZAP70-PLCγ1 (11), Ras-MAPK (12,13), JAK-STAT (14), and PI3K-Akt (15,16). Recently, Andreu-Saumell et al. demonstrated that the intrinsic design of CAR constructs influences their susceptibility to PD-1/PD-L1 inhibition, with high-affinity CARs being more vulnerable to checkpoint suppression, which underscores the importance of checkpoint modulation in optimizing CAR-T cell function (17). As such, disrupting PD-1 signaling has emerged as a promising strategy to enhance CAR-T cell activity.

While several PD-1-targeting strategies are under investigation, variability in efficacy and safety highlights the need for more tunable alternatives such as RNA interference. Although monoclonal antibody-mediated PD-1 blockade can restore CAR-T cells (18,19), this approach is often limited by systemic toxicity (20), heterogeneous patient responses (21,22), and the potential to inadvertently promote tumor growth through interactions with PD-1 expressed on tumor cells (23,24). While CRISPR/Cas9-mediated PD-1 knockout has demonstrated high antitumor efficacy in preclinical models (25,26), complete deletion of PD-1 in naïve CD8⁺ T cells has also been associated with increased exhaustion and impaired cell survival and function (27). In light of these limitations, short hairpin RNA (shRNA)-mediated knockdown offers a more tunable and less disruptive approach to modulate PD-1 expression in CAR-T cells. Liu et al. reported improved anti-tumor activities in mesothelin-targeted CAR-T cells following PD-1 silencing (28). Similarly, Zhou et al. demonstrated that shRNA-mediated PD-1 knockdown augmented CAR-T efficacy across prostate cancer and leukemia xenograft models (29). Clinically, Ma et al. documented successful application of PD-1-knockdown CLL-1 CAR-T cells in two acute myeloid leukemia (AML) patients with post-transplant relapse (30).

Despite recent advances, the optimal shRNA sequence for PD-1 knockdown and its effects on CD19 CAR-T cell exhaustion, memory phenotype, long-term antitumor function, and safety profile have not been systematically evaluated. We hypothesize that PD-1 knockdown can enhance CD19 CAR-T cell persistence, thereby improving their efficacy in PD-L1⁺ tumor environments by alleviating exhaustion and preserving memory subsets. In this study, we engineered CD19-specific CAR-T cells incorporating shRNA-mediated PD-1 knockdown and screened for the most efficient shRNA sequence. We then systematically assessed their phenotype and functional capacity under both repeated and non-repeated antigen stimulation in vitro, as well as evaluated their cytotoxicity and safety in vivo. We present this article in accordance with the MDAR and ARRIVE reporting checklists (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-938/rc).

Methods

DNA constructs and lentivirus production

The third-generation CD19-specific CAR plasmid was a gift from Wuhan University of Science and Technology. The CAR backbone consists of a CD8 hinge, a CD28 transmembrane domain, a CD28 intracellular costimulatory domain, a 4-1BB intracellular costimulatory domain and a CD3ζ intracellular activation domain (31) (Figure 1A). The sequence of FMC63 scFv come from previous patent (patent number: WO 2022/123613 A1).

Figure 1 Screening and validation of shRNAs targeting PD-1. (A) Schematic representation of CD19-CAR and shPD-1-CD19-CAR construct. (B) PD-1 expression in JURKAT cells following CD3/CD28 activation (n=3 independent wells). (C) PD-1 expression on human CD3⁺ T cells after CD3/CD28 stimulation (n=3 donors). (D) PD-1 knockdown efficiency in human CD3⁺ T cells transduced with candidate shRNAs (n=3 donors). (E) Representative flow cytometry plots of PD-1 expression on CAR-T cells after antigen-specific activation. (F) Quantification of PD-1+% in CAR-T cells following antigen stimulation (n=4 donors). (G) PD-1 mRNA expression in CAR-T cells at 24 and 72 h after antigen stimulation, as determined by qPCR (one representative donor from three is shown; n=3 independent wells). Data are presented as mean ± SD. One-way (F) or two-way (G) ANOVA followed by Šídák’s multiple comparisons test was used. **, P<0.01; ***, P<0.001. ANOVA, analysis of variance; CAR, chimeric antigen receptor; CD19, cluster of differentiation 19; PD-1, programmed cell death protein 1; SD, standard deviation; shRNA, short hairpin RNA.

We inserted the shRNA and H1 promoter using molecular cloning technology (Table 1). The sequence structure was as follows: XbaI + sense + loop + antisense + stop signal + Eco31I. Single-stranded DNA oligonucleotide were synthesized and annealed to form double strands. To produce CD19, or shPD-1-CD19 CAR lentivirus, 293T (Human Embryonic Kidney 293T) cells were transfected with a combination of plasmids containing PTK-CAR, pMDLg-pRRE, pRSV-rev, and pMD 2. G.

Cell lines and cell culture

The K562 (human chronic myeloid leukemia, TCHu 191), RAJI (human Burkitt’s lymphoma, TCHu 44) and JURKAT (acute T-cell leukemia cell line, TCHu 96) cell lines were purchased from the Cell Bank of the Chinese Academy of Sciences (CASCB, CSTR: 19375.09.3101HUMTCHu44 and CSTR: 19375.09.3101HUMTCHu191, respectively; Shanghai, China) and cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS). Both cell lines were maintained in a humidified incubator at 37 ℃ with 5% CO₂. The Lenti-X 293T (transformed human embryonic kidney cell line) cell line was obtained from TAKARA (catalog No. 632180) and cultured in DMEM medium with 10% FBS under the same conditions. All culture media and FBS were sourced from Gibco.

To generate RAJI cell line derivatives, a lentiviral vector was constructed to express PD-L1 or both PD-L1 and Luciferase under the control of the Eukaryotic translation elongation factor 1 alpha (EF-1α) promoter with a puromycin selection marker. RAJI cells were transduced with the lentivirus to establish the RAJI-PD-L1 and RAJI-PD-L1-LUCI cell lines. These cells were maintained in RPMI 1640 medium supplemented with 10% FBS and puromycin.

Human peripheral blood mononuclear cells (hPBMCs) from healthy donors were isolated by Ficoll density gradient centrifugation, followed by positive selection of T lymphocytes using CD3-specific magnetic beads. The isolated T cells were stimulated for 24 hours using TransAct reagent and subsequently transduced with lentiviral vectors at a multiplicity of infection (MOI) of 3 to 5.

T cells or CAR-T cells were cultured in TexMACS™ GMP medium supplemented with interleukin 2 (IL-2) at a final concentration of 1,000 U/mL, HEPES (1 M) at a 1:1,000 dilution, and glutamine at a 1:1,000 dilution. Cell growth was maintained by adding or replacing fresh medium as needed. For medium exchange, cells were centrifuged at 700 ×g for 5 minutes to remove the supernatant, resuspended in fresh medium at a density of 2×10⁶ to 3×10⁶ cells/mL, gently mixed, and then incubated at 37 ℃ in a humidified incubator with 5% CO₂.

Flow cytometry

CAR expression and membrane protein levels were assessed using a Beckman CytoFLEX S flow cytometer. For surface staining, over 5×10⁵ target cells were harvested into 5 mL flow cytometry tubes and washed twice with 1 mL of wash buffer [phosphate-buffered saline (PBS) supplemented with 2% FBS] by centrifugation at 300 ×g for 5 minutes. The supernatant was discarded after each wash. Fluorophore-conjugated antibodies specific to the target proteins were added, and the cells were incubated at 2–8 ℃ for 30 minutes. After staining, cells were washed again, centrifuged, and resuspended in 100 µL of wash buffer. To assess cell viability, 2 µL of 7-AAD Viability Staining Solution was added, followed by a 10-minute incubation at room temperature before flow cytometric analysis.

For apoptosis detection, cells were washed twice with flow cytometry wash buffer and resuspended in 100 µL of 1× binding buffer. Subsequently, 5 µL each of PE-conjugated Annexin V and 7-AAD were added, and the cells were incubated for 10 minutes at room temperature. After staining, 400 µL of 1× binding buffer was added, and samples were immediately analyzed by flow cytometry.

To detect cytokine release, a cytometric bead array (CBA) assay was performed. Briefly, 50 µL of capture beads, 50 µL of the PE Detection Reagent, and 50 µL of mouse serum were added to a 5 mL flow cytometry tube. The mixture was gently vortexed and incubated at room temperature in the dark for 3 hours. After incubation, the samples were washed with 1 mL of CBA wash buffer, centrifuged, and resuspended in 300 µL of CBA wash buffer for acquisition by flow cytometry.

The following antibodies and regents were used: PE anti-human CD279 (PD-1) Antibody (Biolegend, San Diego, CA, USA; 329906), FITC Recombinant Protein L (Biolegend, 303653), APC anti-human CD4 Antibody (Biolegend, 300514), PE/Cyanine7 anti-human CD8 (Biolegend, 980910), Brilliant Violet 421™ anti-human CD45RA Antibody (Biolegend, 304130), PE anti-human CD197 (CCR7) Antibody (Biolegend, 353204), Brilliant Violet 421™ anti-human CD366 (Tim-3) Antibody (Biolegend, 364808), Brilliant Violet 650™ anti-human CD223 (LAG-3) Antibody (Biolegend, 369316), PE anti-human CD274 (B7-H1, PD-L1) Antibody (Biolegend, 329706), APC anti-human CD19 Antibody (Biolegend, 302212), PE Annexin V Apoptosis Detection Kit with 7-AAD (Biolegend, 640934), BD™ CBA Human granulocyte-macrophage colony-stimulating factor (GM-CSF) Flex Set (BD Biosciences, San Jose, CA, USA; 558335), BD™ CBA Human IL-2 Flex Set (BD Biosciences, 558270), BD™ CBA Human interferon-gamma (IFN-γ) Flex Set (BD Bioscience, 560111), BD™ CBA Human TNF Flex Set (BD Bioscience, 560112), BD™ CBA Human IL-6 Flex Set (BD Bioscience, 558276), BD™ CBA Human IL-10 Flex Set (BD Bioscience, 558274).

Quantitative real-time PCR (q-PCR)

Total RNA was extracted from CAR-T cells using the TaKaRa MiniBEST Universal RNA Extraction Kit (Takara Bio Inc., Shiga, Japan) and reverse-transcribed into complementary DNA (cDNA) using TaKaRa Reverse Transcriptase M-MLV (RNase H⁻; Takara Bio Inc.). qPCR was performed on the Applied Biosystems QuantStudio™ 12K Flex system using 2× Roche FastStart Universal Probe Master Mix (Roche Diagnostics, Basel, Switzerland).

TaqMan^® Gene Expression Assays (20×, Life Technologies, Carlsbad, CA, USA) was used for the detection of the following human genes: PD-1 (Hs01550088_m1), CCR7 (Hs01013469_m1), SELL (Hs00174151_m1), TCF7 (Hs01556515_m1), HAVCR2 (Hs00958618_m1), LAG3 (Hs00958444_g1), CTLA4 (Hs00175480_m1), IL2 (Hs00174114_m1), IFNG (Hs00989291_m1), granzyme B (GZMB) (Hs00188051_m1), TNF (Hs00174128_m1), and the reference gene GAPDH (Hs99999905_m1). All probes were labeled with 6FAM™ and MGB. The specific primer and probe sequences of the pre-designed TaqMan^® assays are proprietary and not publicly disclosed.

qPCR was conducted using the following thermal cycling conditions: 95 ℃ for 10 minutes, followed by 40 cycles of 95 ℃ for 15 seconds and 60 ℃ for 1 minute. Gene expression levels were analyzed using the comparative Cq method. The Cq value of each target gene was normalized to GAPDH using the formula: ΔCt = Ct_(target) − Ct_(GAPDH). Relative expression changes were calculated using the 2^−ΔΔCq method: 2^−ΔΔCq = 2^{[−(ΔCq_(sample) − ΔCq_(control))]}.

Calcein AM-based cytotoxicity assay

Target cells were labeled with 5 µM Calcein-AM (Aladdin, Shanghai, China) at 37 ℃ for 30 minutes in a non-CO₂ incubator. After labeling, cells were co-cultured with CAR-T cells at effector-to-target (E:T) ratios of 50:1, 10:1, and 2:1 in 96-well plates for 2.5 hours in a humidified incubator. Following incubation, supernatants were collected, and fluorescence intensity (FI) was measured using a microplate reader (PerkinElmer Victor X3, Waltham, MA, USA) at an excitation wavelength of 485/20 nm and an emission wavelength of 530/25 nm. Corrected percent cytotoxicity was calculated using the following formula: corrected % lysis = (FI cytotoxic T lymphocyte assay − FI spontaneous release)/(FI maximum release − FI spontaneous release).

Cytokine release assay

A cytokine-release assay was performed by co-culturing 5×10⁴ CAR-T cells with RAJI-PD-L1 target cells at an E:T ratio of 10:1 in the absence of exogenous IL-2. After 24 hours of incubation, supernatants were collected, and the concentrations of IFN-γ, IL-2, tumor necrosis factor-alpha (TNF-α), and GZMB were quantified using enzyme-linked immunosorbent assay (ELISA) kits (Cat. VAL104C, VAL110; Bio-Techne China Co., Ltd., Shanghai, China). FI was measured using a microplate reader (PerkinElmer Victor X3).

Proliferation assay

To assess proliferation, CAR-T cells were labeled with CellTrace™ CFSE dye (Invitrogen, Carlsbad, CA, USA; C34554) and co-cultured with RAJI-PD-L1 tumor cells in 6-well plates at an E:T ratio of 1:5. After 3 days of incubation, the percentage of CFSE-diluted T cells was quantified by flow cytometry.

Repeated antigen stimulation assay

CAR-T cells were cultured according to the protocol described above. On day 7, a complete medium exchange was performed. Thereafter, RAJI-PD-L1 target cells were added every 48 hours at an E:T ratio of 1:5. Antigen stimulation was repeated 3 times in total. 48 hours after the final stimulation, CAR-T cells were analyzed for apoptosis, memory T cell subsets, immune checkpoint molecule expression, and cytotoxic function.

Luciferase-based cytotoxicity assay

Luciferase-expressing target cells were seeded into 96-well plates at a density of 1×10⁴ cells per well. CAR-T cells were then added at an E:T ratio of 0.1:1, with a final volume of 200 µL per well. The negative control consisted of target cells cultured without CAR-T cells, with the volume adjusted to 200 µL using culture medium. The positive control was prepared by lysing target cells with 50 µL of 2% Triton X-100. After thorough mixing, plates were incubated at 37 ℃ in a humidified CO₂ incubator for 24 hours. Following incubation, 2 µL of D-luciferin solution was added to each well, gently mixed by pipetting, and incubated for an additional 15 minutes at 37 ℃. Luminescence was measured using a multifunctional microplate reader. Cytotoxicity was calculated using the following formula: Cytotoxicity (%) = 1 − [(Luminescence experimental group − Luminescence positive control)/(Luminescence negative control − Luminescence positive control)] ×100%.

Mouse models

A total of 15 female NOD.Cg-PrkdC^scid Il2rg^tm1/Vst (NPG) mice (8 weeks old, 18–20 g) were purchased from Beijing Vitalstar Biotechnology Co., Ltd. (Beijing, China) and housed at the Laboratory Animal Center of Wuhan University of Science and Technology (Wuhan, China). Mice were maintained in a sterile environment under a 12-hour light/dark cycle at ~23 ℃ and 50% relative humidity, with ad libitum access to food and water. All animal experiments were performed under a project license (No. WKD-Zhang-03) granted by the ethics board of Wuhan University of Science and Technology, in compliance with institutional guidelines for the care and use of animals. A protocol was prepared before the study without registration. Each mouse was intravenously injected with 1×10⁶ RAJI-LUCI cells suspended in 100 µL of PBS (32). Tumor progression was monitored using bioluminescence imaging (BLI). On day 6 post-injection, mice were randomly divided into three groups (n=5 per group) and received intravenous injections of 1×10⁷ CD19 CAR-T cells, shPD-1-CD19 CAR-T cells, or control T cells in 100 µL of PBS. Tumor growth was monitored twice weekly using an in vivo imaging system (PerkinElmer). On day 15, peripheral blood (80 µL) was collected via retro-orbital sampling under anesthesia (3% isoflurane for induction, 1% for maintenance) to assess cytokine release. Tumor radiance curves were randomized basis on initial radiance (before treatment) value to ensure balanced groups. No blinding was applied as they relied on machine-based readout. The health and behavior of the mice were monitored daily. On day 20, or earlier if humane endpoints were met (e.g., >20% body weight loss, persistent hunched posture, severe weakness, inability to eat or drink, labored breathing, or unmanageable tumor burden), mice were euthanized.

Statistical analysis

No statistical methods were used to predetermine sample sizes, but our sample sizes were similar to those reported in previous publications or based on our own experience. Biological replicates represent independent experiments using T cells isolated from at least three different healthy donors. Details on the number of experimental replicates and the nature of the replicates (technical or biological) are specified in the respective figure legends. Statistical analyses were conducted using Prism 9.0 software (GraphPad Software) or Excel. Applied analyses and statistical significances are indicated in the corresponding figures and figure legends. Data were tested for normality by applying the Shapiro-Wilk test. Group comparisons were performed using an unpaired Student’s t-test or analysis of variance (ANOVA) with Šídák’s post hoc multiple comparisons test as appropriate. P value <0.05 was considered statistically significant. Data were presented as the mean ± standard deviation (SD). No datapoints were excluded from analysis.

Results

Identification of a sequence for effective knockdown of PD-1 gene expression

Ten unique shRNA-CD19 CAR plasmids targeting different PD-1 mRNA binding regions were constructed and packaged into lentiviral vectors for transduction into activated JURKAT cells (Figure 1A). Among these, shRNA-3, shRNA-4, shRNA-6, and shRNA-10 demonstrated the most pronounced PD-1 knockdown and were selected for subsequent functional evaluation (Figure 1B).

PD-1 expression was found to be upregulated in activated T cells, reaching a peak on day 3 following CD3/CD28 stimulation before gradually declining (Figure 1C). We therefore evaluated the PD-1 knockdown efficiency of the four selected shRNA constructs at 3-, 4-, and 5-days post-activation. Quantitative analysis revealed that shRNA-3 and shRNA-6 mediated the most significant suppression of PD-1 expression (Figure 1D).

To further evaluate their ability to inhibit tumor-induced PD-1 upregulation, CAR-T cells cultured for 7 days were co-incubated with RAJI target cells at an E:T ratio of 1:5 for 24 hours. Flow cytometric analysis revealed that shPD-1-3-CD19 CAR-T cells showed significantly reduced PD-1 mean fluorescence intensity (MFI) compared to both control CD19 CAR-T cells and shPD-1-6-CD19 CAR-T cells (Figure 1E,1F).

PD-1 transcript levels were subsequently analyzed by quantitative PCR. Total RNA was isolated from CAR-T cells co-cultured with RAJI cells for 24 and 72 hours. Among all groups, shPD-1-3-CD19-CAR-T cells exhibited the most substantial reduction in PD-1 mRNA expression relative to the CD19-CAR-T control (Figure 1G). These findings support the selection of shRNA-3 for subsequent experiments.

PD-1 knockdown relieves tumor-mediated inhibition and enhances the function of CD19-CAR-T cells

Comprehensive phenotypic and functional characterization revealed no significant differences between shPD-1 CD19-CAR-T cells and control CD19 CAR-T cells in terms of proliferative capacity, CD4⁺/CD8⁺ subset distribution, or memory T cell compartment composition (Figure 2A-2E). These findings demonstrate that PD-1 knockdown does not compromise the fundamental properties of CAR-T cells.

Figure 2 The basal characteristics of shPD-1-CD19-CAR and CD19-CAR cells. (A) Fold expansion of CAR-T cells from day 0 to 14, measured by trypan blue exclusion (n=4 donors). (B) Representative flow plots of CD4⁺ and CD8⁺ CAR-T cells. (C) Summarized results of CD4⁺/CD8⁺ ratios of CAR-T cells (n=4 donors). (D) Representative flow plots of the expression of CCR7 and CD45RA on CAR-T cells at day 7. (E) Summarized results of T cell subsets: naïve + TSCM (CCR7⁺CD45RA⁺), TCM (CCR7⁺CD45RA⁻), TEM (CCR7⁻CD45RA⁻), and TEFF (CCR7⁻CD45RA⁺) (n=4 donors). Data are presented as mean ± SD. Unpaired Student’s t-test (C), one-way (E) ANOVA followed by Sidak’s multiple-comparison test is used. ANOVA, analysis of variance; CAR-T, chimeric antigen receptor T cell; CD19, cluster of differentiation 19; ns, not significant; PD-1, programmed cell death protein 1; SD, standard deviation; shRNA, short hairpin RNA; TCM, central memory T cell; TEFF, effector T cell; TEM, effector memory T cell; TSCM, stem cell memory T cells.

To assess antigen-specific functionality, three tumor cell lines were selected: (I) parental RAJI cells (CD19⁺/PD-L1⁻); (II) RAJI-PD-L1 cells (CD19⁺/PD-L1⁺); and (III) K562 cells (CD19⁻/PD-L1⁻) (Figure 3A,3B). shPD-1-CD19 CAR-T and CD19 CAR-T cells displayed comparable cytotoxicity against RAJI cells. However, only shPD-1-modified CAR-T cells exhibited significantly enhanced cytotoxic activity against RAJI-PD-L1 cells, with negligible killing of CD19⁻ K562 cells (Figure 3C). After 24-hour co-culture with target cells, shPD-1 CD19-CAR-T cells secreted higher levels of IL-2, IFN-γ, TNF-α, and GZMB compared with CD19 CAR-T cells both at the protein and transcript levels (Figure 3D,3E). These findings demonstrate that genetic PD-1 blockade selectively enhances CAR-T cell effector function in PD-L1⁺ tumor microenvironments.

Figure 3 PD-1 knockdown relieves tumor-mediated inhibition and enhances the function of CD19-CAR-T cells. (A) CD19 expression on RAJI and K562 cell lines. (B) PD-L1 expression on RAJI, RAJI-PD-L1, and K562 cells. (C) Cytotoxicity of CAR-T cells against tumor cell lines at different E:T ratios (2:1, 10:1, 50:1), measured by Calcein-AM release assay after 2.5 h of co-culture (one representative donor from three is shown; n=4 independent wells). (D) Cytokine secretion after 24 h co-culture with RAJI-PD-L1 cells (one representative donor from three is shown; n=3 independent wells). (E) qPCR analysis of cytolytic gene expression relative to GAPDH after 24 h co-culture (one representative donor from three is shown; n = 3 independent wells). (F) CFSE dilution in CAR-T cells after 72 h co-culture with RAJI-PD-L1 cells (n=4 donors). (G) Annexin V/7-AAD staining of CAR-T cells following 72 h co-culture. (H) Quantification of apoptotic CAR-T cells (n=4 donors). Data are presented as mean ± SD. Unpaired Student’s t-test (E,H), one-way (D,F) or two-way (C) ANOVA followed by Sidak’s multiple-comparison test (D) is used. *, P<0.05; **, P<0.01; ***, P<0.001; ****, P<0.0001; ns, not significant. AAD, 7-aminoactinomycin D staining; ANOVA, analysis of variance; CAR-T, chimeric antigen receptor T cell; CD19, cluster of differentiation 19; CFSE, carboxyfluorescein succinimidyl ester; E:T, effector-to-target; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GZMB, granzyme B; IFN-γ, interferon-gamma; IL-2, interleukin 2; PD-1, programmed cell death protein 1; PD-L1, programmed death ligand 1; SD, standard deviation; TNF-α, tumor necrosis factor-alpha.

To evaluate the effect of PD-1 knockdown on cellular proliferation and viability, we performed CFSE-based proliferation assays following 72-hour co-culture with PD-L1-expressing RAJI cells. shPD-1-CD19-CAR-T cells exhibited significantly greater CFSE dilution compared to controls (Figure 3F). Concurrently, Annexin V/7-AAD staining revealed a lower apoptosis rate in the shPD-1-CD19 CAR-T group compared to controls (Figure 3G,3H). These indicate that PD-1 knockdown simultaneously enhanced proliferation and reduced activation-induced cell death (AICD), thereby conferring a survival advantage.

PD-1 knockdown preserves memory phenotypes and reduces exhaustion following repeated antigen stimulation

The therapeutic efficacy of CAR-T cells is closely linked to their persistence (33). To investigate whether PD-1 knockdown enhances CAR-T cell durability under chronic antigen exposure, a repeated stimulation model was established. CAR-T cells were co-cultured with RAJI-PD-L1 cells at an E:T ratio of 1:5, with stimulation repeated every 48 hours for 3 cycles (Figure 4A). After the 3rd stimulation, shPD-1-CD19 CAR-T cells exhibited significantly lower apoptosis rates than the CD19 CAR-T group (Figure 4B,4C).

Figure 4 PD-1 knockdown maintains memory phenotype and reduces exhaustion after repeated antigen stimulation. (A) Schematic of in vitro repeated antigen stimulation (“stress test”) model. CAR-T cells were challenged every 48 h with RAJI-PD-L1 cells at a 1:5 E:T ratio. (B) Representative flow plots of Annexin V/7-AAD staining after repeated antigen stimulation rounds. (C) Apoptosis rates of CAR-T cells after repeated antigen exposure (n=4 donors). (D) Representative CCR7 and CD45RA expression after repeated stimulation. (E) Summarized results of memory T cell subsets: TN + TSCM (CCR7⁺CD45RA⁺), TCM, TEM, and TEFF (n=4 donors) after repeated stimulation. (F) Expression of memory-related genes after repeated stimulation rounds (one representative donor from three is shown; n=3 independent wells). (G) Representative flow plot of the expression of exhaustion markers (e.g., TIM-3, LAG-3) on CAR-T cells after repeated stimulation. (H) Percentages of TIM-3⁺ and LAG-3⁺ CAR-T cells after repeated stimulation (n=4 donors). (I) Expression of exhaustion-related genes after repeated stimulation rounds (one representative donor from three is shown; n=3 independent wells). (J) Cytotoxic activity of CAR-T cells after 3 rounds of antigen stimulation (one representative donor from three is shown; n=4 independent wells). Data are presented as mean ± SD. Unpaired Student’s t-test (C,E,H), two-way ANOVA followed by Sidak’s multiple-comparison test (F,I,J) is used. *, P<0.05, **, P<0.01; ***, P<0.001; ****, P<0.0001. AAD, 7-aminoactinomycin D staining; ANOVA, analysis of variance; CAR-T, chimeric antigen receptor T cell; CD19, cluster of differentiation 19; E:T, effector-to-target; PD-1, programmed cell death protein 1; PD-L1, programmed death ligand 1; SD, standard deviation; TCM, central memory T cell; TEFF, effector T cell; TEM, effector memory T cell; TN, naïve T cells; TSCM, stem cell memory T cells.

Flow cytometric analysis revealed that the shPD-1-CD19 CAR-T group showed a significantly higher proportion of memory T cell subsets, including naïve T cells (TN) and stem cell memory T cells (TSCM) (Figure 4D,4E). Consistently, qPCR demonstrated upregulation of CCR7, SELL (encoding CD62L), and TCF7, which are known regulators of memory phenotype maintenance and lymphoid tissue homing (34,35) (Figure 4F).

Exhaustion marker analysis showed a reduction in T cell immunoglobulin and mucin domain-containing protein 3 (TIM-3)⁺ and Lymphocyte activation gene-3 (LAG-3)⁺ populations in shPD-1-CD19-CAR-T cells compared to controls (Figure 4G,4H), supported by decreased mRNA levels of HAVCR2, LAG3, and CTLA4 (Figure 4I). These findings suggest that PD-1 knockdown not only counteracts PD-1/PD-L1-mediated suppression but also attenuates the expression of multiple inhibitory receptors.

To assess CAR-T cell functionality, cytotoxicity assays were performed after three rounds of antigen stimulation. Although both CAR-T groups showed reduced cytotoxicity compared to unstimulated levels, only shPD-1-CD19 CAR-T cells retained measurable killing activity against target cells, whereas the control CD19-CAR-T cells completely lost cytolytic function (Figure 4J).

PD-1 knockdown enhances antitumor efficacy of CD19-CAR-T cells in vivo

To assess therapeutic efficacy in vivo, NPG mice were intravenously injected with RAJI-PD-L1-LUCI tumor cells. 7 days post-injection, mice were treated with 1×10⁷ shPD-1-CD19-CAR-T cells, CD19-CAR-T cells (control group), or control T cells (control group) via the tail vein (Figure 5A). BLI revealed that shPD-1-CD19-CAR-T cells persistently inhibited tumor progression, whereas rapid tumor progression was observed in both the control T cell and unmodified CD19 CAR-T cell groups (Figure 5B,5C). By day 19 post-treatment, the survival duration of mice treated with shPD-1-CD19 CAR T cells was prolonged compared to those treated with CD19 CAR T cells or control T cells (Figure 5D).

Figure 5 PD-1 knockdown improves antitumor efficacy of CD19-CAR-T cells in vivo. (A) Experimental timeline of xenograft model using RAJI-PD-L1-LUCI cells. (B) Tumor burden over time assessed by bioluminescence imaging. (C) Quantification of tumor burden (total flux, photons/s) (n=5 mice). (D) Kaplan-Meier survival curves of mice treated with CD19-CAR-T, shPD-1-CD19-CAR-T, or control T cells (n=5 mice). (E) Serum cytokine levels measured by CBA assay (n=4–5 mice). Data are presented as mean ± SD. Unpaired Student’s t-test is used. *, P<0.05; **, P<0.01; ns, not significant. CAR, chimeric antigen receptor; CBA, cytometric bead array; CD19, cluster of differentiation 19; GM-CSF, granulocyte-macrophage colony-stimulating factor; IFN-γ, interferon-gamma; IL-2, interleukin 2; NPG, NOD.Cg-PrkdC^scid Il2rg^tm1/Vst; PD-1, programmed cell death protein 1; PD-L1, programmed death ligand 1; TNF-α, tumor necrosis factor-alpha.

We next analyzed cytokine release profiles in peripheral blood. The shPD-1-modified CAR-T cells secreted significantly higher levels of IFN-γ, TNF-α, and GM-CSF compared to unmodified CAR-T cells. Notably, IL-6 levels—a key mediator of cytokine release syndrome (CRS)—remained at comparable levels between groups (Figure 5E), suggesting that PD-1 knockdown augments CAR-T cell effector function without exacerbating CRS-related toxicity.

Discussion

T cell exhaustion, characterized by the progressive loss of effector functions and sustained expression of inhibitory receptors, is a major barrier to effective immunotherapy (36-39). A key driver of this dysfunctional state is the immunosuppressive tumor microenvironment, which induces persistent upregulation of immune checkpoint molecules—particularly PD-1—on T cells through chronic antigen stimulation and suppressive signaling, ultimately leading to functional impairment. In this study, we developed a shRNA-based strategy to downregulate PD-1 expression in CD19 CAR-T cells. Systematic screening identified an optimal shRNA sequence that achieved robust knockdown in CD19 CAR-T cells. This modification significantly enhanced CAR-T cell antitumor activity both in vitro and in vivo. Notably, PD-1 knockdown preserved memory T cell phenotypes and maintained cytotoxicity following repeated antigen stimulation, simulating chronic tumor exposure. Furthermore, enhanced cytokine production, likely due to alleviation of exhaustion, contributed to superior tumor control in a PD-L1⁺ xenograft model, without evidence of increased CRS risk.

We performed a systematic screening of multiple shRNA sequences targeting PD-1, aiming to identify the most effective construct for stable and functional gene silencing in CAR-T cells. Candidate sequences included several shRNAs previously reported in the literature (28,40). The most effective shRNA was subsequently incorporated into the CD19 CAR construct to generate shPD-1-CD19 CAR-T cells for further functional and phenotypic analyses. Importantly, this shRNA not only reduced PD-1 expression under resting conditions, but also significantly suppressed PD-1 upregulation following CD19 antigen-mediated CAR activation—a key mechanism contributing to CAR-T cell exhaustion in the tumor microenvironment.

Consistent with previous studies linking memory T cell phenotype to improved clinical outcomes (41,42), The repeated antigen stimulation assay mimics the tumor microenvironment by exposing CAR-T cells to persistent tumor antigen challenge. This in vitro model revealed that shPD-1-CD19 CAR-T cells exhibited improved persistence and effector function under chronic stimulation, which may explain their superior antitumor activity observed in vivo. Meanwhile, we also observed that upregulated TCF7 expression on shPD-1-CD19 CAR-T cell after repeated antigen stimulation assays, a transcription factor critical for Wnt/β-catenin signaling and the regulation of CCR7 and SELL—key mediators of lymphoid homing and memory maintenance (34,35). These findings provide further evidence supporting the link between PD-1 and Wnt/β-catenin signaling, highlighting the necessity for further investigation to confirm the mechanistic connections between them.

Additionally, we observed significantly reduced expression levels of TIM-3 and LAG-3 in shPD-1-CD19 CAR-T cells compared to control CAR-T cells following repeated stimulation. One possible explanation is that early relief of PD-1-mediated inhibition delays the onset of terminal exhaustion, thereby reducing upregulation of checkpoints such as TIM-3 and LAG-3. The reduction of these inhibitory signals may further diminish the exhausted phenotype of CAR-T cells. This sequential downregulation of exhaustion markers may establish a reinforcing loop that sustains CAR-T functionality in suppressive environments.

CRS is a potentially life-threatening toxicity of CAR-T cell therapy, largely mediated by IL-6, which serves as both a key driver and clinical biomarker of this condition (43). Immune checkpoint inhibition, including PD-1 blockade, can amplify T cell activation and has been associated with an increased risk of CRS (44). However, in our study, shPD-1-CAR-T cells showed comparable IL-6 levels to control CAR-T cells, despite elevated production of IFN-γ and TNF-α. This may be attributed to the cell-intrinsic nature of PD-1 gene silencing, which avoids systemic immune activation commonly observed with antibody-based checkpoint blockade. Nevertheless, further safety profiling, including broader cytokine analysis and long-term in vivo toxicity studies, will be essential for clinical translation.

Our findings suggest that the PD-1 knockdown strategy holds strong translational potential for clinical application in CD19 CAR-T cell therapy. Functionally, PD-1 knockdown enhanced CAR-T cell persistence and long-term antitumor efficacy both in vitro and in vivo. From a technical perspective, shRNA-mediated checkpoint modulation can be seamlessly incorporated into existing CAR-T manufacturing workflows without substantially impacting production timelines or regulatory pathways. Moreover, this approach may be extended to solid tumors, where PD-1/PD-L1-mediated immunosuppression remains a significant barrier to effective therapy. Nonetheless, further clinical studies are needed to evaluate the safety, efficacy, and feasibility of this strategy in patients.

Conclusions

These findings indicate that PD-1-targeting shRNA can alleviate the immunosuppressive effects on human anti-CD19 CAR-T cells, thereby representing a promising strategy to enhance their therapeutic efficacy and translational potential.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the MDAR and ARRIVE reporting checklists. Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-938/rc

Data Sharing Statement: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-938/dss

Peer Review File: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-938/prf

Funding: This research was supported by the 2023 Hubei Provincial Key Research and Development Plan (No. 2023BCB090); and 2019 Hubei Province Technology Innovation Special Major Project (No. 2019ACA168).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-938/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All animal experiments were performed under a project license (No. WKD-Zhang-03) granted by the ethics board of Wuhan university of science and technology, in compliance with institutional guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

References

1. Zugasti I, Espinosa-Aroca L, Fidyt K, et al. CAR-T cell therapy for cancer: current challenges and future directions. Signal Transduct Target Ther 2025;10:210. [Crossref] [PubMed]
2. Dabas P, Danda A. Revolutionizing cancer treatment: a comprehensive review of CAR-T cell therapy. Med Oncol 2023;40:275. [Crossref] [PubMed]
3. Velasco Cárdenas RM, Brandl SM, Meléndez AV, et al. Harnessing CD3 diversity to optimize CAR T cells. Nat Immunol 2023;24:2135-49. [Crossref] [PubMed]
4. Grover P, Veilleux O, Tian L, et al. Chimeric antigen receptor T-cell therapy in adults with B-cell acute lymphoblastic leukemia. Blood Adv 2022;6:1608-18. [Crossref] [PubMed]
5. Gardner RA, Shah NN. CAR T-Cells for Cure in Pediatric B-ALL. J Clin Oncol 2023;41:1646-8. [Crossref] [PubMed]
6. Jacoby E, Bielorai B, Hutt D, et al. Parameters of long-term response with CD28-based CD19 chimaeric antigen receptor-modified T cells in children and young adults with B-acute lymphoblastic leukaemia. Br J Haematol 2022;197:475-81. [Crossref] [PubMed]
7. Cappell KM, Sherry RM, Yang JC, et al. Long-Term Follow-Up of Anti-CD19 Chimeric Antigen Receptor T-Cell Therapy. J Clin Oncol 2020;38:3805-15. [Crossref] [PubMed]
8. Poorebrahim M, Melief J, Pico de Coaña Y, et al. Counteracting CAR T cell dysfunction. Oncogene 2021;40:421-35. [Crossref] [PubMed]
9. Cheng J, Xiao Y, Peng T, et al. ETV7 limits the antiviral and antitumor efficacy of CD8(+) T cells by diverting their fate toward exhaustion. Nat Cancer 2025;6:338-56. [Crossref] [PubMed]
10. Sailer CJ, Hong Y, Dahal A, et al. PD-1(Hi) CAR-T cells provide superior protection against solid tumors. Front Immunol 2023;14:1187850. [Crossref] [PubMed]
11. Parvez A, Choudhary F, Mudgal P, et al. PD-1 and PD-L1: architects of immune symphony and immunotherapy breakthroughs in cancer treatment. Front Immunol 2023;14:1296341. [Crossref] [PubMed]
12. Liotti F, Kumar N, Prevete N, et al. PD-1 blockade delays tumor growth by inhibiting an intrinsic SHP2/Ras/MAPK signalling in thyroid cancer cells. J Exp Clin Cancer Res 2021;40:22. [Crossref] [PubMed]
13. Jeon SM, Lim JS, Park SH, et al. Blockade of PD-L1/PD-1 signaling promotes osteo-/odontogenic differentiation through Ras activation. Int J Oral Sci 2022;14:18. [Crossref] [PubMed]
14. Syamsu SA, Faruk M, Smaradania N, et al. PD-1/PD-L1 pathway: Current research in breast cancer. Breast Dis 2024;43:79-92. [Crossref] [PubMed]
15. Quan Z, Yang Y, Zheng H, et al. Clinical implications of the interaction between PD-1/PD-L1 and PI3K/AKT/mTOR pathway in progression and treatment of non-small cell lung cancer. J Cancer 2022;13:3434-43. [Crossref] [PubMed]
16. Wang S, Liu C, Yang C, et al. PI3K/AKT/mTOR and PD‑1/CTLA‑4/CD28 pathways as key targets of cancer immunotherapy Oncol Lett 2024;28:567. (Review). [Crossref] [PubMed]
17. Andreu-Saumell I, Rodriguez-Garcia A, Mühlgrabner V, et al. CAR affinity modulates the sensitivity of CAR-T cells to PD-1/PD-L1-mediated inhibition. Nat Commun 2024;15:3552. [Crossref] [PubMed]
18. Gray KD, McCloskey JE, Vedvyas Y, et al. PD1 Blockade Enhances ICAM1-Directed CAR T Therapeutic Efficacy in Advanced Thyroid Cancer. Clin Cancer Res 2020;26:6003-16. [Crossref] [PubMed]
19. Cillo AR, Cardello C, Shan F, et al. Blockade of LAG-3 and PD-1 leads to co-expression of cytotoxic and exhaustion gene modules in CD8(+) T cells to promote antitumor immunity. Cell 2024;187:4373-4388.e15. [Crossref] [PubMed]
20. Wang SJ, Dougan SK, Dougan M. Immune mechanisms of toxicity from checkpoint inhibitors. Trends Cancer 2023;9:543-53. [Crossref] [PubMed]
21. Zurko J, Chaney K, Astle JM, et al. PD-1 blockade after bispecific LV20.19 CAR T modulates CAR T-cell immunophenotype without meaningful clinical response. Haematologica 2021;106:2788-90. [Crossref] [PubMed]
22. Mu J, Deng H, Lyu C, et al. Efficacy of programmed cell death 1 inhibitor maintenance therapy after combined treatment with programmed cell death 1 inhibitors and anti-CD19-chimeric antigen receptor T cells in patients with relapsed/refractory diffuse large B-cell lymphoma and high tumor burden. Hematol Oncol 2023;41:275-84. [Crossref] [PubMed]
23. Yao H, Wang H, Li C, et al. Cancer Cell-Intrinsic PD-1 and Implications in Combinatorial Immunotherapy. Front Immunol 2018;9:1774. [Crossref] [PubMed]
24. Du S, McCall N, Park K, et al. Blockade of Tumor-Expressed PD-1 promotes lung cancer growth. Oncoimmunology 2018;7:e1408747. [Crossref] [PubMed]
25. Zhang J, Hu Y, Yang J, et al. Non-viral, specifically targeted CAR-T cells achieve high safety and efficacy in B-NHL. Nature 2022;609:369-74. [Crossref] [PubMed]
26. Choi BD, Yu X, Castano AP, et al. CRISPR-Cas9 disruption of PD-1 enhances activity of universal EGFRvIII CAR T cells in a preclinical model of human glioblastoma. J Immunother Cancer 2019;7:304. [Crossref] [PubMed]
27. Odorizzi PM, Pauken KE, Paley MA, et al. Genetic absence of PD-1 promotes accumulation of terminally differentiated exhausted CD8+ T cells. J Exp Med 2015;212:1125-37. [Crossref] [PubMed]
28. Liu G, Zhang Q, Li D, et al. PD-1 silencing improves anti-tumor activities of human mesothelin-targeted CAR T cells. Hum Immunol 2021;82:130-8. [Crossref] [PubMed]
29. Zhou JE, Yu J, Wang Y, et al. ShRNA-mediated silencing of PD-1 augments the efficacy of chimeric antigen receptor T cells on subcutaneous prostate and leukemia xenograft. Biomed Pharmacother 2021;137:111339. [Crossref] [PubMed]
30. Ma YJ, Dai HP, Cui QY, et al. Successful application of PD-1 knockdown CLL-1 CAR-T therapy in two AML patients with post-transplant relapse and failure of anti-CD38 CAR-T cell treatment. Am J Cancer Res 2022;12:615-21.
31. Shi J, Zhang Z, Cen H, et al. CAR T cells targeting CD99 as an approach to eradicate T-cell acute lymphoblastic leukemia without normal blood cells toxicity. J Hematol Oncol 2021;14:162. [Crossref] [PubMed]
32. Li D, Qin J, Zhou T, et al. Bispecific GPC3/PD‑1 CAR‑T cells for the treatment of HCC. Int J Oncol 2023;62:53. [Crossref] [PubMed]
33. Wittibschlager V, Bacher U, Seipel K, et al. CAR T-Cell Persistence Correlates with Improved Outcome in Patients with B-Cell Lymphoma. Int J Mol Sci 2023;24:5688. [Crossref] [PubMed]
34. Escobar G, Mangani D, Anderson AC. T cell factor 1: A master regulator of the T cell response in disease. Sci Immunol 2020;5:eabb9726. [Crossref] [PubMed]
35. Szabo PA, Levitin HM, Miron M, et al. Single-cell transcriptomics of human T cells reveals tissue and activation signatures in health and disease. Nat Commun 2019;10:4706. [Crossref] [PubMed]
36. Kouro T, Himuro H, Sasada T. Exhaustion of CAR T cells: potential causes and solutions. J Transl Med 2022;20:239. [Crossref] [PubMed]
37. Ahn T, Bae EA, Seo H. Decoding and overcoming T cell exhaustion: Epigenetic and transcriptional dynamics in CAR-T cells against solid tumors. Mol Ther 2024;32:1617-27. [Crossref] [PubMed]
38. Minton K. Overcoming CAR T cell exhaustion. Nat Rev Immunol 2020;20:72-3. [Crossref] [PubMed]
39. Zhu X, Li Q, Zhu X. Mechanisms of CAR T cell exhaustion and current counteraction strategies. Front Cell Dev Biol 2022;10:1034257. [Crossref] [PubMed]
40. Lee YH, Lee HJ, Kim HC, et al. PD-1 and TIGIT downregulation distinctly affect the effector and early memory phenotypes of CD19-targeting CAR T cells. Mol Ther 2022;30:579-92. [Crossref] [PubMed]
41. Baysal MA, Chakraborty A, Tsimberidou AM. Enhancing the Efficacy of CAR-T Cell Therapy: A Comprehensive Exploration of Cellular Strategies and Molecular Dynamics. J Cancer Immunol (Wilmington) 2024;6:20-8. [Crossref] [PubMed]
42. Tao Z, Chyra Z, Kotulová J, et al. Impact of T cell characteristics on CAR-T cell therapy in hematological malignancies. Blood Cancer J 2024;14:213. [Crossref] [PubMed]
43. Kang S, Tanaka T, Inoue H, et al. IL-6 trans-signaling induces plasminogen activator inhibitor-1 from vascular endothelial cells in cytokine release syndrome. Proc Natl Acad Sci U S A 2020;117:22351-6. [Crossref] [PubMed]
44. Zhang X, You Y, Zhang P, et al. Cytokine release syndrome caused by immune checkpoint inhibitors: a case report and literature review. Future Sci OA 2024;10:2422786. [Crossref] [PubMed]