Peripheral blood mononuclear cells (PBMCs) were obtained from healthy patient samples in accordance with our Stony Brook University Institutional Review Board (IRB) approved protocol. All donors provided written informed consent. The CCRF-CEM cell line was acquired from ATCC (Manassas VA). T cells and CCRF-CEM cells were cultured as previously described (23, 24).

The lentiviral vector was generated as previously described (23, 25–27). HEK293T cells were cultured in T flasks to achieve approximately 70-80% confluence. Next, cells were transfected with the CD7-RTX CAR containing expression plasmid and viral packaging plasmid as previously described (23). Incubation was performed as previously described (23). Following incubation, the resulting supernatant was harvested, filtered through a 0.2 µM disc filter, and stored at -80°C until utilized.

The activation and transduction procedures of T cells utilizing lentiviral vector was described previously (23). CD7-RTX CAR T cell expression was analyzed using flow cytometry to detect F(ab)’ fragment, following previously described protocol (23). Flow cytometry was conducted using an FACS Calibur instrument (Becton Dickinson, Franklin Lakes, NJ), and results were analyzed using Kaluza software (Beckman Coulter, CA).

CD7-RTX CAR T cells or control T cells were incubated with CD7 expressing CCRF-CEM cells in 18 hour 1:1 or 2:1 co-culture experiments in 1 mL of T cell growth media, excluding IL-2, as previously described (23). Following 18 hours of co-culture, cells were stained with mouse anti-human CD3 (Cytek Tonbo) and CD7 (BD Biosciences) and then analyzed by flow cytometry.

In vivo activity of the CD7-RTX CAR T cell therapy was assessed in NOD scid gamma (NSG) mice that were sub-lethally irradiated and intravenously injected with luciferase-expressing CCRF-CEM cells. This study used all male mice, 8 to 12 weeks of age, with an approximate weight range of 25-30 grams obtained from Jackson Laboratories (Bar Harbor, ME, USA), strain (NOD.Cg-PrkdcsidIl2rgtm1Wjl/SzJ). All animal experimentation was performed in accordance with Stony Brook University’s Department of Laboratory Resources (DLAR), under an approved IACUC protocol in accordance with good laboratory practice (GLP) guidelines.

Depletion of the CD7-RTX CAR T therapy through infusion of rituximab was verified in vivo. Each mouse was infused with 10×10⁶ CD7-RTX CAR T cells (Day 1) and subsequently injected with 150 µg (5 mg/kg) of rituximab on Days 5, 6, 7, 9 and 13. On Day 15, all mice were sacrificed for whole blood analysis. Blood samples were labeled using CD3, CD45, CD34 and F(ab)’ antibodies.

The manufacturing of CAR T cells was carried out in a good manufacturing practices (GMP) laboratory. Peripheral blood apheresis PBMCs were obtained from allogeneic donors (the patient’s son, HLA 6/10 matched). After isolation of PBMCs, T cells were activated and transduced with lentiviral vector CD7-RTX for 48 hours. Subsequently, CAR cells were cultured and then tested for the presence of pathogenic microorganisms and contaminants. The CAR T cells were labeled and analyzed by flow cytometry to determine transduction efficiency.

The patient provided written informed consent for publication. A 53-year-old male first presented to the hospital in February 2018 with an erythematous, pruritic, disseminated rash over the total body surface. The patient was initially treated for allergic dermatitis, however, his condition progressed. In April 2019, biopsies of the left axillary lymph node and skin as well as peripheral blood analysis confirmed the diagnosis of SS with peripheral blood, bone marrow and lymph node involvement. With reference to the International Society for Cutaneous Lymphomas and European Organization of Research and Treatment of Cancer (ISCL/EORTC) staging system, the disease was categorized as T2N3B1M0 IVA-2 (28). After 1.5 years of palliative care, the malignancy progressed with symptomatic worsening of skin and lymph node disease. In May 2021, routine blood testing demonstrated significant leukocytosis: white blood cells (WBC) 41×10⁹/L, neutrophils (NEU) 11×10⁹/L (additional values: hemoglobin (HGB) 130g/L, platelets (PLT) 455x10⁹/L). Peripheral blood and bone marrow morphology indicated abnormal T lymphocytes. The bone marrow contained 53.76% of nuclear cells and expressed surface antigens CD3, TCRα/β, CD7, CD2, CD4, CD45RO, and weakly expressed CD5. The following surface markers were not expressed: CD8, CD56, CD57, CD30, CD16, CD45RA, CD26, CD10, CD25, granzyme B, perforin, CDL161, CD94, CD38, and TCRγ/δ. Bone marrow biopsy demonstrated clonal T cell proliferation, bone marrow invasion, as well as positive TCRγ and TCRβ rearrangement, negative TCD rearrangement, and ATM gene negative. The patient was assessed for lymphoma associated gene mutations which yielded positive results for TET2, BCORL1, KMT2B, PLCG2, TYK2 mutations and TP53 gene deletion. The skin pathological T cell phenotype was CD3(+), CD5(+), CD7(+), CD4(+, partial), CD8(-), PD-1(-). From May to October 2021, 8 courses of GVD (Gemcitabine, Vinorelbine, and Pegylated Liposomal Doxorubicin) and Dexamethasone were administered (first line of therapy), which led to symptom improvement, rash reduction by 60%, and lymph node size reduction by 50% indicating partial response (PR). Stable disease was confirmed by peripheral blood testing. Complications were observed following first-line treatment, notably, pulmonary bacterial and fungal infections, sacral and caudal skin infections (roseomonas mucosa and staphylococcus lugdunensis), as well as deep vein thrombosis. In November 2021, the patient returned with worsening lymph node enlargement, in addition to overall disease progression. Three courses of Brentuximab Vedotin, Chidamide (a HDACi) and thalidomide chemotherapy were administered as second line therapy, which led to stabilization of the disease. In February 2022, 3 courses of Mitoxantrone Liposomes combined with Selinexor were administered as third line therapy, which also resulted in disease stabilization (SD). However, severe allergic reaction appeared after the administration of Selinexor. In June 2022, a skin biopsy taken from the abdomen and the thigh indicated 10% tumor cell volume with a high tumor burden. After which, the patient was administered Venetoclax and Homoharringtine (HHT, an alkaloid derived from trees of the genus Cephalotaxus), Decitabine and Chidamide, Thalidomide, Methotrexate, and Interferon as fourth line therapy, and subsequently Camrelizumab (a PD-1 inhibitor) as fifth line therapy. The patient’s condition proved to be refractory to several lines of therapy consistent with disease relapse. No therapy trialed achieved and maintained complete response (CR). Given the patient’s severe r/r disease and CD7 surface antigen expression, the patient became eligible for treatment with our CD7-RTX CAR T therapy.

The expression of our second generation CD7-RTX CAR T therapy is driven by a spleen focus-forming virus (SFFV) promoter. The extracellular CAR components are comprised of an anti-CD7 single-chain variable fragment (scFv), CD8-derived hinge (H) and transmembrane (TM) domains as well as a “safety switch” RTX domain. The intracellular domains consist of CD28 and CD3ζ signaling moieties ( Figure 1A ).

The CD7-RTX CAR T therapy is engineered through a single step, non-gene editing approach whereby the CAR binds to and internalizes the CD7 surface antigen, thus avoiding fratricide. Flow cytometry analysis demonstrated approximately 72-81% of T cells expressed the CD7-RTX CAR and that greater than 99% of T cells did not express CD7 ( Figures 1B–D ). Despite the loss of CD7 surface expression, CD7-RTX CAR T cells demonstrated comparable expansion to control T cells ( Figure 1E ).

CD7-RTX T cells were co-cultured with CCRF-CEM cells and incubated for 18 hours. Potent lysis of CCRF-CEM cells was confirmed as the CD7-RTX CAR T cells achieved 99.88% and 99.82% depletion, relative to controls, at 1:1 and 2:1, respectively ( Figure 1F ).

To establish a xenograft mouse model, 12 NSG mice received a sub-lethal dose of gamma irradiation (2.0 Gy) and were assigned randomly to treatment or control groups. 1.0×10⁶ CCRF-CEM cells were infused 24 hours post-irradiation.

Five days following introduction of CCRF-CEM cells, the mice were intravenously injected with 10×10⁶ CD7-RTX CAR or control T cells. To evaluate tumor burden on days 5, 10, 13, 16, and 19, RediJect D-Luciferin (Perkin-Elmer) was injected intraperitoneally and the mice were subjected to IVIS imaging to quantify luciferase activity. While flux, and thus tumor burden, continuously increased in control mice, it remained at approximately background levels for those treated with CD7-RTX CAR T cells ( Figures 2A, B ). While control mice were observed to have significant residual tumor burden in peripheral blood analysis, mice treated with CD7-RTX CAR T cells demonstrated remarkable >99% complete depletion ( Figures 2A, C ). Additionally, mice infused with the CD7-RTX CAR T therapy showed significant survival improvement (p=0.0026) in comparison to control T cell-treated mice ( Figure 2D ).

The CD7-RTX CAR T therapy demonstrated sustained absence of the CD7 surface antigen from T cells in vivo. Peripheral blood was extracted from the mice on day 20 and day 43, 15 days post-transduction and 38 days post-infusion, respectively, and evaluated by flow cytometry. Cells were first plotted against CD45 and side scatter (SSC). The CD45⁺ cells were then gated and analyzed for surface expression of CD3 and CD7 antigens. At day 20, CD7-RTX CAR T treated mice were observed to have approximately >94% CD3⁺/CD7^- T cells, demonstrating sustained downregulation of the CD7 surface antigen ( Supplementary Figure 1A ). By day 43, approximately >99% CD3⁺/CD7^- cells were observed ( Supplementary Figure 1B ).

The depletion of CD7-RTX CAR T cells through the infusion of rituximab was validated in vivo. Six NSG mice were sub-lethally irradiated and intravenously injected with 10×10⁶ CD7-RTX CAR T cells. Following CAR T cell expansion, on day 5, three mice were randomly injected with saline (control) or rituximab (150µg). Mice who received rituximab demonstrated approximately 96% reduction of CD7-RTX CAR T cells in peripheral blood as compared to controls ( Supplementary Figure 1C ).

A patient suffering from r/r SS was treated as compassionate use with CD7-RTX CAR T cells. Prior to CAR T treatment, advanced disease status was confirmed in February 2023, characterized by lymph node infiltration, extensive cutaneous disease, and presence in peripheral blood. On physical examination, the patient displayed an extensive full body, pruritic, erythematous skin rash ( Figure 3A ). Before treatment, malignant T-lineage cells were quantified at 67.83% ( Figure 3D , upper panels) by flow cytometry, and demonstrated a single peak, suggesting clonal origin, by T cell receptor (TCR) evaluation ( Figure 3D , upper panel). Histopathological analysis demonstrated a substantial burden of CD7⁺ malignant cells within the skin prior to CAR-treatment ( Figure 3C ).

Fludarabine and cyclophosphamide were administered for conditioning prior to CD7-RTX CAR T cell therapy (February 3, 2023). The patient was infused with about 2×10⁶ CD7-RTX CAR T cells/kg body weight (February 8, 2023). Evaluation of peripheral blood one-week post-CD7-RTX CAR T treatment revealed partial remission (PR) with 97.29% reduction in tumor burden. Peripheral blood assessment at day 12 indicated >99% reduction in tumor burden with molecular complete remission (CR) achieved at day 28 post-treatment ( Figures 3D, E ), and resolution of cutaneous disease ( Figure 3B ). Day 28 post-CAR infusion, TCR morphology results indicated polyclonal T-lymphocyte presence with a greater than 3-fold reduction in maximum peak intensity ( Figure 3D , bottom panel). No signs of graft versus host disease (GVHD) were found during the follow-up period.

Positron emission tomography-computed tomography (PET-CT) imaging was performed prior to treatment and demonstrated high uptake in lesions over the right temple, right axilla, and left inguinal region ( Figure 4A ). Repeat imaging at 1 month, 10 months, 16 months post-infusion demonstrated absence of signal suggesting resolution of disease ( Figures 4B–D ).

The patient underwent haploidentical allogeneic hematopoietic stem cell transplantation (the donor was his son, the same donor of CAR T cells) about one month after CD7-RTX CART treatment. The last follow-up date was September 2024, which was 18 months after bone marrow transplant, and the patient was in continuous CR.

Peripheral blood levels of interleukin (IL)-2 and interferon (INF)-γ demonstrated a decrease from baseline eight days (D8) following CD7-RTX CAR T treatment, and both returned to D0 levels, 15 days post-treatment ( Figures 5A, B ). Analysis of IL-6 indicated a rapid increase in the cytokine (50-60 pg/mL) approximately 1-2 days post-infusion which returned to near undetectable levels by D7 ( Figure 5C ). Ferritin levels remained near constant throughout, trending between 150-200 pg/mL from D0 to D22 ( Figure 5D ). The maximum cytokine levels and the time of peak (days post-CAR) can be found in Supplementary Table 1 . Following infusion of CD7-RTX T cells the percentage of CD7⁺ malignant T cells diminished rapidly as compared to the total PBMC count ( Figure 5E ). Expansion of CD7 negative RTX CAR T cells was observed as levels increased from D5 (50%) to D8 (100%), before returning on D12 (25%) and remaining approximately constant to D28 ( Figure 5F ). CD8⁺ T cell expansion was observed in proportion to CD4⁺ T cells ( Figure 5G ), corresponding to the infusion and replication of CD7-RTX T cells on D8 ( Figure 5F ). On day 28, the patient’s peripheral blood T cell count had recovered to the normal range (1834 cells/µL), whereas the NK cell count, although gradually increasing to 65.97 cells/µL, remained below the normal threshold (77 cells/µL) and exhibited fluctuations. ( Figures 5H, I ).

Remarkable safety was demonstrated with no severe adverse events (SAEs). The patient was found to have pneumonia post-treatment, consistent prior pneumonia history, however this resolved with supportive care. Two days following CD7-RTX CAR T cell treatment the patient developed a fever of 39.2°C and tachycardia with a maximum heart rate of 120 beats per minute (BPM), both of which resolved with supportive care. No other adverse events (AEs) occurred.