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. 2025 Jul 19;176(4):520–529. doi: 10.1111/imm.70018

Use of a Foamy‐Virus Vector System to Produce an ‘Off‐the‐Shelf’ Fcγ‐CR‐T Cell Product for the Treatment of Haematological and Solid Tumour Malignancies

Ioanna Lazana 1,2,✉, Emmanouil Simantirakis 1, Evangelos Kourous 1, Maria Daniil 1, Dimitris Ioannou 1, George Vassilopoulos 1,3
PMCID: PMC12583232  PMID: 40682568

ABSTRACT

The emergence of chimeric antigen receptor (CAR)‐T cells has revolutionised the therapeutic landscape of hematologic malignancies, with limited translation only to solid organ malignancies. Furthermore, safety concerns have recently been raised, in terms of tumorigenesis, relating to the viral vector systems used for gene transduction, underlying the need for alternative, safer gene therapy delivery systems. In this study, we investigated the use of foamy virus (FV) vectors, known for their favourable integration profile and reduced genotoxicity, to generate Fcγ (CD16)‐chimeric receptor (CR) T cells. Our aim was to provide T cells with the capacity to recognise and bind to tumour cells, opsonised with commercially available monoclonal antibodies (mAbs), enhancing the antibody‐dependent cell cytotoxicity (ADCC). The high‐affinity F158V polymorphism of the CD16, previously shown to mediate superior ADCC, was utilised for the generation of Fcγ (CD16)‐CR T cells. Results showed that FV‐derived CD16‐CR T cells exhibited robust CD16 expression and demonstrated potent functional activity, including: (i) high mAb‐binding capacity, (ii) formation of T‐cell tumour cell aggregates in the presence of mAbs, (iii) significant degranulation and proinflammatory cytokine production upon tumour engagement and (iv) potent, dose‐dependent cytotoxic activity against target cells in the presence of mAbs. This is the first study, to our knowledge, demonstrating that FV vectors can be used to generate potent CD16‐CR T cells, providing a safer and more versatile platform for antibody‐based immunotherapy. This approach enables the expansion of engineered T cell therapies beyond hematologic malignancies and towards solid tumours, using clinically approved mAbs.

Keywords: cancer, cell therapy, Fc receptors, immunotherapy

Despite the huge success of chimeric antigen receptor (CAR)‐T cells in the treatment of hematologic malignancies, safety concerns have recently been raised in terms of tumorigenesis, relating to the viral vector systems used for product generation, underlying the need for alternative methods for gene modification. Our group has generated a safe, ‘universal’, immune cell therapy, with potent cytotoxic activity, for the treatment not only of haematological but also of solid organ malignancies, by providing healthy T cells with the capacity to bind tumour cells, in the presence of commercially available monoclonal antibodies, supplementing their anti‐tumour toxicity.

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1. Introduction

The emergence of chimeric antigen receptor (CAR)‐T cell therapies has transformed the therapeutic landscape of haematological malignancies [1, 2]. A chimeric receptor (CR) is a half B‐cell and half T‐cell receptor; the B‐part can recognise a specific tumour antigen and the T‐part can trigger toxicity. When such receptors are introduced into immune cells (such as T cells), the cells get re‐directed to kill tumour cells bearing the specific antigen [3, 4]. It is also of particular importance to note that tumour antigen recognition by the CAR occurs through the B‐cell receptor and thus it bypasses the restrictions of the HLA‐mediated antigen recognition posed by the TCR. This allows the use of an individual's T‐cells to generate CAR‐T cells for tumour eradication. Another advantage of CAR‐T cell therapies is the provision of an improved immunogenic memory leading to a continuous immune surveillance that can prevent and/or treat future relapses [5].

Despite the huge success of such therapies in haematological malignancies, there has been poor translation to solid organ malignancies [6]. This is mainly attributed to the following reasons: (i) the lack of unique tumour‐associated antigens, (ii) the poor penetration of cytotoxic cells to tumours and, most importantly, (iii) the immunosuppressive microenvironment, rendering cytotoxic cells into anergic cells [7, 8]. Initial attempts to use first generation CAR‐T cells, targeting different tumour antigens (such as anhydrase IX, folate receptor‐a, human epidermal growth factor receptor 2 [HER2], fibroblast activation protein [FAP], etc.) have been associated with disappointing results due to the on‐target/off‐tumour effects, poor persistence (due to anti‐CAR responses), limited CAR‐T cell homing and poor anti‐tumour activity [9].

The emergence of monoclonal antibodies (mAbs) targeting specific tumour antigens, such as anti‐CD20 (Rituximab) and anti‐EGFR (Cetuximab), has revolutionised the landscape of cancer therapeutics. Their tumoricidal action is mainly mediated through antibody‐dependent cell‐mediated cytotoxicity (ADCC), which is exerted through the Fc receptor of immune cells, such as NK cells, γδ‐T cells, neutrophils and macrophages [10]. However, their clinical efficacy gradually declines with disease progression, owing to a decrease in the number of active immune effector cells and/or to their exhaustion [11, 12]. Furthermore, polymorphisms of the Fc gamma receptor (FcγR) genes (having either a phenylalanine, F, or a valine, V, residue at position 158) have been suggested to greatly impact the response to antibody treatment, owing to variable binding capacity of the Fc receptor [13]. More specifically, the F158V polymorphism has been associated with a higher Fc binding, resulting in higher tumoricidal activity and better patient outcomes [14].

CD16 is an FcγR that binds IgG molecules through their Fc portion. Given the critical role of CD16‐expressing immune effector cells in ADCC, we generated Fcγ‐CR‐T cells, recognising the Fc part of mAbs used for the treatment of malignancies. More specifically, we gene‐modified T cells from healthy individuals to express the Fcγ‐158 V receptor in order to be used as an adoptive cell therapy for ADCC, in conjunction with commercially available mAbs. Similar efforts had been made before by various groups, using lentiviral (LV) vectors to introduce the FcγR, with encouraging results [13, 14, 15]. LV vectors have been widely used for gene modification, including for the generation of CD16‐CR T cells [16]. However, safety concerns on the use of LV vectors have recently emerged, with one study reporting the development of myelodysplastic syndrome (MDS)/acute myeloid leukemia (AML) in a patient who underwent gene therapy as a cure for sickle cell disease [17] and another one revealing transduction of a leukaemic blast with an anti‐CD19 CAR in a patient with B‐acute lymphoblastic leukaemia (ALL), leading to relapse and death [18]. In 2022, the FDA issued a warning about the risk of developing T cell malignancies after the infusion of virally transduced CAR T‐cell products [19, 20]. Following this report, further cases of T cell malignancies have been reported. It is of note that in some cases, a causality analysis indicated that the malignancy was related to vector insertion [21, 22, 23]. This has led to the investigation of alternative strategies for gene modification. Foamy virus (FV) vectors are considered to be a safe alternative, owing to their unique characteristics: (i) broad tissue tropism, (ii) safe integration profile, (iii) large transgene capacity and (iv) less read‐through transcription [24, 25, 26]. Given their favourable safety profile, we explored the use of FV vectors for the generation of allogeneic CD16‐F158V‐CR T cells, with the aim to be used as a single, ‘universal’, cell therapy for the treatment of malignancies for which there is a commercially available mAb.

2. Materials and Methods

2.1. Cell Lines

HEK‐293T cells (Lenti‐X 293T; Takara Clontech, ECACC Cat #12022001, RRID:CVCL_0063), used for viral vector production, were cultured in DMEM (Sigma‐Aldrich, Germany) supplemented with 10% heat‐inactivated fetal calf serum (FCS; Gibco) and 100 U/mL penicillin–streptomycin (P/S) and maintained at 37°C in a humidified 5% CO2 atmosphere. EGFR+ human cancer cell lines, Panc‐1 (pancreatic adenocarcinoma; ECACC, RRID:CVCL_0480) and DLD‐1 (colorectal adenocarcinoma; ATCC, RRID:SCR_001672) and the CD19+ B cell lymphoma line Raji (ATCC, RRID:CVCL_0511) were cultured under identical conditions (Panc‐1, DLD‐1 in DMEM; Raji in RPMI 1640; Lonza, Switzerland).

2.2. Human Samples and T Cell Cultures

Peripheral blood samples were obtained from healthy donors with informed consent, under approval from the Institutional Review Board of the Biomedical Research Foundation of the Academy of Athens, in accordance with the Declaration of Helsinki. Peripheral blood mononuclear cells (PBMCs) were isolated by density gradient centrifugation (Histopaque‐1077; Sigma‐Aldrich) and T cells were purified using a negative selection magnetic bead kit (Dynabeads Untouched Human T Cells Kit; Invitrogen). T cells were stimulated with anti‐CD3/CD28 beads (TransAct; Miltenyi Biotec) and cultured in Optimizer medium (Gibco) supplemented with 2.5% T‐cell expansion supplement, 5% FCS, 1% l‐glutamine and 100 U/mL P/S. IL‐7/IL‐15 (10 ng/mL; PeproTech) was added and replenished every 2–3 days.

2.3. Plasmid Construction

A codon‐optimised CD16‐F158V CR was synthesised (ITD) and cloned into FV and LV backbones. The construct included a CD8α leader sequence, the CD16‐F158V extracellular domain and intracellular CD28 and CD3ζ domains.

For FV vector generation, the CR16 cassette was subcloned into pDF‐MCS using Gibson Assembly. The vector backbone was linearised with BamHI and NotI (NEB). PCR amplification of the CR16 insert used specific primers (FVEF1F and FVCR‐R) and KAPA HiFi polymerase. Assembly products were transformed into NEBstable Escherichia coli , and colonies screened by PCR and verified via Sanger sequencing (Eurofins).

For LV vectors, the CR16 cassette was PCR‐amplified using primers (LVC16R and LVEF) and inserted into EcoRV/SalI‐digested pRRLSIN.cPPT.PGK‐GFP.WPRE (Addgene; gift from D. Trono). Correct assembly was confirmed as above (details of the construction of FV‐ and LV‐viral vectors are documented in Data S1).

2.4. Viral Production and T Cell Transduction

FV particles were produced by co‐transfecting HEK‐293 T cells with 24 μg FV CR16 plasmid and 13.5 μg packaging plasmids (PCIN/GS, PCIN/PS, PCIN/ES) using Lipofectamine 2000 (Invitrogen). LV particles were generated using 20 μg LV CR16 and 13 μg packaging plasmids (psPAX2 and pMDG2). Viral supernatants were harvested 24–72 h post‐transfection, filtered (0.45 μm) and concentrated by ultracentrifugation at 27 000g for 90 min.

For viral titration, 105 HEK 293‐T cells were transduced with various concentrations of the isolated FV and LV viral stocks for 24 and 72 h, respectively. At the end of the incubation period, the % expression of CD16 was assessed by flow cytometry. The virus titre was calculated based on the following formula: Virus titre = % expression (×) No. of cells used/virus volume.

Activated T cells (106 cells/mL) were transduced with FV or LV particles (MOI 5–10 for FV, 10–20 for LV) in the presence of heparin or polybrene, respectively. Transductions were enhanced by centrifugation at 1200g for 90 min at 32°C. Post‐transduction, cells were expanded in Optimizer medium containing 10 ng/mL IL‐7 and 10 ng/mL IL‐15 (PeproTech).

2.5. Antibodies and Reagents

The following antibodies were used: CD16‐BV421, CD3‐FITC, CD4‐BV786, CD8‐BV421 (BD Biosciences) and 7AAD (Cayman Chemical, USA). Flow cytometry was conducted using a BD FACS Celesta Cell Analyser and data analysis was performed using FlowJo Version 7.2.2 Software (RRID:SCR_008520). Rituximab and Cetuximab were a kind gift from the Department of Pharmacy, Larisa University Hospital, Greece.

2.6. Antibody Binding, Cell Aggregation

To measure the CR's antibody‐binding capacity, CR‐T cells were incubated with Rituximab (0.1 μg/mL) or Cetuximab (0.1 μg/mL) for 30 min and stained with PE‐conjugated goat anti‐human IgG (BD Biosciences). In order to identify the optimal mAb concentration, a kinetic was performed using increasing concentrations (0.01, 0.05, 0.1, 0.5 and 1 μg/mL) of mAbs.

To determine whether the addition of mAb enables the binding between CD16‐CR T cells and tumour cells, CD16‐stained CR‐T cells were incubated with either Rituximab or Cetuximab and mixed with CFSE‐labelled target cells (Raji, DLD and Panc‐1) at a 1:1 ratio. The formation of aggregates (defined as double positive—CFSE and CD16—cells) was assessed by flow cytometry. The reverse was also assessed: CFSE‐labelled target tumour cells were incubated with mAbs and mixed with CD16‐stained CR‐T cells at a ratio of 1:1.

2.7. Cell Activation and Degranulation Assays

CD16‐CR T cells were mixed with Raji, DLD1 or Panc01 cells in the presence or absence of mAbs (Rituximab or Cetuximab) for 48 h. Culture supernatants were assessed for IFN‐γ levels by ELISA (ThermoFisher Scientific).

To determine whether antibody binding to the CR promoted degranulation, CD16‐stained FV‐ or LV‐derived Fcγ‐CR‐T cells were mixed with target cells, with or without mAbs. The CD107a‐PE antibody (BD Biosciences) was added to the culture and the % CD107a positive cells were assessed by flow cytometry.

2.8. Cytotoxicity

Fcγ‐CR T‐cells were incubated with target cells (Raji, DLD or Panc‐1) at a ratio of 5:1 and 10:1 for 18 h, in the presence or absence of mAbs (Rituximab or Cetuximab). The number of viable cells (CFSE positive/7AAD negative cells) was assessed by flow cytometry and the % cytotoxicity was calculated as follows: [1 − live targets (sample)/live targets (control)] × 100.

2.9. Statistical Analysis

Data were analysed using GraphPad Prism Version 5.0 (GraphPad Software Inc., San Diego, CA, USA, RRID:SCR_002798). Student's t test, Mann–Whitney or Fisher's exact test were carried out according to the nature of the data. p < 0.05 was considered significant.

3. Results

3.1. Generation of CD16‐CR Construct and Expression of the CD16 Chimeric Receptor

The 158V polymorphism of the CD16 gene is associated with an increased Fc‐terminus antibody affinity, leading to better responses to antibody therapy [27, 28, 29]. A CR gene construct was generated, consisting of the CD16‐158V binding domain, the transmembrane region of CD8 and the intracellular CD28 domain in tandem with the CD3ζ signalling molecule.

FV is considered to be a safer viral vector, having a favourable integration site profile, than the commonly used LV vectors [25]. For that reason, FV gene‐modified CD16‐CR T cells (FV‐CD16‐CR T cells) were generated, the efficacy of which was compared to that of LV gene‐modified CD16‐CR T cells (LV‐CD16‐CR T cells). CD16 expression levels were found to be comparable between the FV‐CD16‐CR T cells and the LV‐CD16‐CR T cells (mean ± SEM, 74.6% ± 4.1% vs. 81.2% ± 3.9%, respectively, p = 0.3), (Figure 1A). However, it has to be noted that a considerably lower MOI was used for the generation of FV‐CD16‐CR T cells (MOI 5–10), compared to that used for the generation of LV‐CD16‐CR T cells (MOI 10–20). Fold expansion at Day 14 was higher, although not significantly, in the FV—compared to the LV‐CD16‐CR T cells (mean ± SEM, 3.1 ± 0.4 vs. 2.8 ± 0.6 × 102‐fold increase, p = 0.09), (Figure 1B). The CD8 T‐cell population expanded in both FV‐ and LV‐CD16‐CR T cells by Day 14 (mean ± SEM, 64.1% ± 1.4% vs. 61.9% ± 2.3%, p = 0.13), (Figure 1C).

FIGURE 1.

FIGURE 1

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Production and expansion of foamy virus (FV)‐derived CD16‐CR T cells. (A) Transduction efficiency of FV‐ and lentiviral (LV)‐derived CD16‐CR T cells. The graph illustrates the CD16 expression levels of T cells transduced with FV and LV vectors. (B) Fold expansion of FV‐ and LV‐derived CD16‐CR T cells. Cells were expanded in ex vivo cultures supplemented with 10 μg/mL human recombinant IL7 and 10 μg/mL human recombinant IL‐15, as illustrated (n = 3). Fold expansion was calculated by dividing the number of cells on Days 7 and 14 by the number of cells on Day 0. (C) Percentage (%) ratio of CD4/CD8 cells during the cell culture and expansion of FV‐ and LV‐derived CD16‐CR T cells (n = 3). Error bars indicate SEM.

3.2. CD16‐CR T Cells Bind Effectively Monoclonal Antibodies

In order to assess the binding capacity of the CD16‐CR, FV‐ and LV‐CD16‐CR T cells were incubated with the clinically available mAbs Rituximab (anti‐CD20) and Cetuximab (anti‐EGFR). A concentration of 0.1 μg/mL was used, based on a kinetic analysis using various mAb concentrations (Figure S1).

The mean antibody binding capacity of the FV‐CD16‐CR T cells was 68.7% ± 3.7% and 71.3% ± 4.1% for Rituximab and Cetuximab, respectively, whereas that of LV‐CD16‐CR T cells was 72.1% ± 6.2% and 76.5% ± 7.4% for Rituximab and Cetuximab, respectively. No significant difference was noted between the two (p = 0.17, n = 3), (Figure 2).

FIGURE 2.

FIGURE 2

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Monoclonal antibody (mAb)‐binding capacity of FV‐ and LV‐transduced CD16‐CR T cells. Untransduced T‐cells are unable to bind mAbs, as opposed to FV‐ and LV‐derived CD16‐CR T cells, which bind strongly the commercially available mAbs, Rituximab and Cetuximab. A representative experiment is illustrated (A), as well as a summary graph of three experiments (B). CR; chimeric receptor; FV; foamy virus, IgG; immunoglobulin G; LV; lentivirus.

3.3. Monoclonal Antibodies as a Bridge Between CD16‐CR T Cells and Tumour Cells

To assess whether the mAb‐conjugated FV‐ and LV‐CD16‐CR T cells are able to bind to tumour cells, aggregation assays were performed. More specifically, the FV‐CD16‐CR and LV‐CD16‐CR T cells were incubated with the mAb Rituximab or Cetuximab. Following this initial binding step, the cells were mixed with CFSE‐labelled target cells (Raji, DLD and Panc01 cells) and the formation of aggregates was assessed by flow cytometry. Results showed that only in the presence of mAbs the FV‐ and LV‐CD16‐CR T cells were able to bind to target cells (Figure 3).

FIGURE 3.

FIGURE 3

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Aggregation studies. The graph demonstrates the monoclonal antibody (mAb)‐mediated binding of CD16‐CR T cells to tumour cells. As the graph shows, untransduced T cells (A) do not bind tumour cells, irrespective of the presence of mAb, whereas FV‐ and LV‐derived CD16‐CR T cells (B) come to proximity and bind tumour cells, forming aggregates, only in the presence of mAbs. (C) Summary of aggregation assays (mean ± SD of three experiments). As illustrated, significantly higher % cell aggregation was noted when CD16‐CR T cells (both FV‐ and LV‐derived), as opposed to untransduced T cells, were combined with target cells (Raji, DLD or Panc01 cells), in the presence of mAbs. FV, foamy virus; Ig, immunoglobulin; LV, lenti virus.

When the target cells were incubated with mAb (Rituximab or Cetuximab), as an initial step, before they were incubated with the FV‐ and LV‐CD16‐CR T cells, similar results were obtained, indicating that the mAbs act as a bridge between the CD16‐CR T cells and the tumour cells, irrespective of the timing of mAb administration. This is of particular importance for the potential clinical use of the CD16‐CR T cells, as the mAb may be administered either prior to the CD16‐CR T cell infusion (saturating tumour cells) or at the same time (incubating CD16‐CR T cells with mAbs prior to their administration).

3.4. Monoclonal Binding to CD16‐CR T Cells Induces Cell Activation and Degranulation

Having confirmed the mAb‐binding capacity of the FV‐ and LV‐CD16‐CR T cells, we investigated whether the mAb‐mediated recognition of tumour cells by CD16‐CR T cells promoted cell activation, with subsequent degranulation. For that reason, CD16‐stained FV‐ or LV‐CD16‐CR T cells were incubated with target cells (Raji, DLD and Panc01 cells), in the presence or absence of mAb. A significant increase in the production of IFN‐γ, a proinflammatory cytokine, was noted in the presence of mAbs, as compared to cells (target and effectors) alone. This increase was comparable between the FV‐ and LV‐CD16‐CR T cells, as illustrated in Figure 4A.

FIGURE 4.

FIGURE 4

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Monoclonal antibody binding to CD16‐CR T cells induces cell activation and degranulation. (A) FV‐ and LV‐derived CD16‐CR T cell (depicted as ‘+’) production of the proinflammatory cytokine IFN‐γ, in response to mAb (either Rituximab or Cetuximab)‐binding of tumour cells (Raji and DLD1 and Panc01 cells, respectively), as determined by ELISA. Error bars represent SDs. Non‐gene modified T cells are depicted as ‘−’. Degranulation of (B) FV‐ and (C) LV‐derived CD16‐CR T cells induced by the addition of Abs (Rituximab and Cetuximab) to cancer cells (Raji and DLD1 and Panc01, respectively). A representative result from three experiments, using different donors for the generation of CD16‐CR T cells is shown. (D and E) Degranulation data are shown across two different effector to target ratios (1:1 and 5:1). Significantly increased % degranulation of FV‐ (D) and LV‐derived (E) CD16‐CR T cells is noted with the addition of mAbs (depicted as ‘+’) at both ratios.

Furthermore, the addition of mAb (both Rituximab and Cetuximab) promoted a significant degree of degranulation, as compared to cells alone. More specifically, the mean % degranulation of FV‐CD16‐CR cells was as follows: (i) 39.8% (SEM, 3.2%) with the addition of Rituximab to Raji cells, (ii) 29.1% (SEM, 4.5%) with the addition of Cetuximab to DLD1 cells and (iii) 23.3% (SEM, 1.9%) with the addition of Cetuximab to Panc01 cells (Figure 4B). Again, the addition of mAbs to LV‐CD16‐CR T cells exhibited comparable levels of degranulation. Overall, these results demonstrate the functional activation of CD16‐CR T cells (both FV‐ and LV‐modified) upon binding of the mAb.

3.5. Monoclonal Antibodies Induce Potent ADCC Activity of CD16‐CR T Cells

The cytotoxic activity of the FV‐ and LV‐CD16‐CR T cells was assessed, after incubation with CFSE‐labelled target cells (Raji, DLD and Panc01 cells), at a ratio of 5:1 and 10:1, in the presence or absence of mAb (Rituximab 0.1 μg/mL or Cetuximab 0.1 μg/mL, respectively). Results demonstrated that the addition of mAbs significantly enhanced the cytolytic activity of both the FV‐ and the LV‐CD16‐CR T cells in a dose‐dependent way. More specifically, the mean % cytotoxicity of FV‐CD16‐CR T cells was: (ii) 36.8% and 51.5% at 5:1 and 10:1 ratio, respectively, in the presence of Rituximab and Raji cells, (ii) 46.5% and 57.7% at 5:1 and 10:1 ratio, respectively, in the presence of Cetuximab and DLD1 cells and (iii) 31.4% and 40.3% at 5:1 and 10:1 ratio, respectively, in the presence of Cetuximab and Panc01 cells (Figure 5). For LV‐CD16‐CRs, the respective % lysis was comparable. These results support a potent mAb‐induced ADCC activity of FV‐CD16‐CR T cells, comparable to that of LV‐CD16‐CR T cells.

FIGURE 5.

FIGURE 5

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Cytotoxicity assays: (A) The graph illustrates the % cytotoxicity of FV‐ and LV‐derived CD16‐CR T cells against Raji, DLD and Panc01 cells, in the presence or absence of a monoclonal antibody (mAb: Rituximab or Cetuximab, respectively), in an effector to target ratio (E:T) of 5:1 and 10:1 (n = 3). Significantly increased % cytotoxicity is demonstrated with the addition of mAbs, indicating that CD16‐CR T cells are able to induce a potent mAb‐dependent tumour cell toxicity The bars depict the mean with standard deviation. (B) A representative experiment out of three is illustrated. Again, a significant % cell toxicity is demonstrated in the presence of mAbs, in a dose‐dependent way.

4. Discussion

The clinical success of CAR‐T cell therapy has transformed the treatment landscape for relapsed/refractory haematological malignancies, offering durable remissions to patients with otherwise poor prognosis [30, 31, 32]. However, despite substantial efforts, this success has not been replicated in solid tumours. Barriers such as antigen heterogeneity and the poor penetration of cells into the tumour microenvironment contribute to this disparity [6, 33, 34, 35]. To address these challenges, alternative strategies, such as the generation of Fcγ (CD16)‐CR T cells, have been explored by various groups [14, 15, 36]. These cells are engineered to recognise mAb‐opsonised tumour cells, enhancing the antibody‐dependent cellular cytotoxicity (ADCC) [37]. A key advantage of CD16‐CR T cells is their modularity; they can be redirected towards diverse tumour antigens simply by co‐administering the appropriate clinically approved mAb [38]. Additional benefits may include reduced off‐target toxicity (through mAb withdrawal), the potential to mitigate acute toxicity (through saturation of CD16‐CR T cells by immunoglobulins) and most importantly, the potential to simultaneously target multiple antigens [36, 39].

Recent safety concerns surrounding integrating viral vectors, particularly the risk of insertional mutagenesis and secondary T cell malignancies, have renewed the interest in safer vector platforms. In 2022, the FDA issued warnings regarding these events in the context of CAR‐T cell therapy. Although the incidence of virally induced secondary malignancies remains low, there is a pressing need for novel strategies to be developed, minimising the risk of secondary primary malignancies. To this end, we evaluated the feasibility and functionality of a FV vector for CD16‐CR T cell generation. FV vectors are considered promising for clinical application due to their: (i) broad tissue tropism, (ii) favourable integration profile, (iii) large transgene capacity and (iv) reduced likelihood of read‐through transcription [40, 41, 42]. This is the first study, to our knowledge, describing the successful generation of CD16‐CR T cells, using FV vectors. All experiments were conducted in parallel with LV vectors to establish functional comparability.

It is well known that the affinity of the Fcγ receptor for the Fc part of an antibody plays a crucial role in ADCC, influencing clinical responses to antibody immunotherapy [43]. In our study, the F158V polymorphism of the CD16 gene was selected, given the higher binding capacity of the encoded receptor, mediating superior ADCC [13, 29, 44, 45, 46]. The CR design incorporated CD3ζ and CD28 signalling domains to provide both activation and co‐stimulatory signals, in line with second‐generation receptor constructs shown to enhance T cell expansion and function [47, 48].

FV‐CD16‐CR T cells exhibited high transduction efficiency, comparable to LV‐CD16‐CR T cells, but at a lower MOI (5–10 vs. 10–20, respectively). This is of particular importance for clinical translation, as lower MOIs reduce vector use and consequently manufacturing costs and production time and minimise the cytotoxicity related to excessive viral exposure. The resulting CD16 expression levels were consistent with prior reports on LV‐based constructs [11, 39].

We then explored the antibody‐binding capacity of the CD16‐CR T cells, by incubating the cells with commercially available mAbs (such as Rituximab and Cetuximab). Results showed a high binding capacity of both FV‐ and LV‐derived CD16‐CR T cells, at comparable levels. The ability of mAb to act as a bridge between the CD16‐CR T cells and the tumour cells was assessed in two ways. First, we incubated CD16‐CR T cells with mAbs, following which the ‘coated’ cells were incubated with tumour cells and the % aggregation was assessed. Results showed that only in the presence of mAbs there was aggregate formation (between CD16‐CR T cells and tumour cells), whereas no such binding was noted in the absence of mAb. Similar results were noted when CD16‐CR T cells were incubated with mAb‐coated tumour cells. The later finding has been previously reported by other groups [15, 39]. However, the fact that mAb‐pre‐coated CD16‐CR T cells bind equally effectively to the tumour cells open new horizons in CD16‐CR therapy, as this would potentially abrogate any potential side effects associated with the non‐specific binding of the CD16‐CR T cells in vivo. This approach may help mitigate risks in contexts such as autoimmune disorders or IgG‐mediated pathologies, where tissue‐bound antibodies might inadvertently engage CD16‐CR T cells [14].

Then, we investigated the potential stimulation of CD16‐CR T cells and subsequent proinflammatory cytokine production and cell degranulation upon binding with mAbs and tumour cells. Results revealed that CD16‐CR T cells produced significant levels of IFN‐γ upon mAb‐dependent recognition of target cells, consistent with T cell activation. Degranulation was also significantly enhanced, underscoring the capacity of FV‐CD16‐CR T cells to mediate potent ADCC. These results are consistent with previous studies suggesting that the conventional cytolytic activity may be complemented by granule‐independent mechanisms in tumour eradication [39]. Finally, cytotoxicity assays further demonstrated that both FV‐ and LV‐CD16‐CR T cells efficiently lysed EGFR+ and CD19+ tumour cells in the presence of their respective mAbs, in a dose‐dependent manner. The observed tumour cell toxicity was in line with prior reports on CD16‐CR T cells engineered using LV vectors [38, 39].

To conclude, this is the first study, to our knowledge, providing evidence that FV vectors can be used to generate functional CD16‐CR T cells, capable of mediating potent mAb‐dependent anti‐tumour responses, comparable to their LV‐engineered counterparts. Overall, our data add to the evidence that adoptive cell therapy, using CD16‐CR T cells, may enhance ADCC and boost immune responses by promoting antibody‐dependent T cell responses. This is of particular clinical importance, as it allows for a more versatile immune therapy, which: (i) supplements the use of the currently available commercial mAbs, (ii) allows a potential use of multiple mAbs with the same CD16‐CR T cell therapy, targeting different tumour antigens, (iii) has a more manageable side effect profile (reduction of CD16‐CR T cell toxicity, by removal of the mAb or administration of high doses of immunoglobulins) and (iv) carries a minimal risk of vector‐induced tumorigenesis. Although further in vivo studies are warranted to validate safety and efficacy, our data provide a strong foundation for continued translational development of a safer and more versatile adoptive immunotherapy, such as the FV‐CD16‐CR T cell therapy.

Ethics Statement

This study was performed in accordance with the Declaration of Helsinki and the protocol was approved by the Local Ethics Committee (Approval No: EHΔΕ7/18‐09‐2022).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1. The binding capacity of various concentrations of the monoclonal antibodies (mAbs) Rituximab and Cetuximab to FV‐derived CD16‐CR T cells is illustrated. A dose‐dependent binding was noted, which starts to plateau at 0.1 μg/mL. This concentration (0.1 μg/mL) has been used in all subsequent experiments. μg/mL, micrograms per millilitre.

IMM-176-520-s001.docx (517KB, docx)

Data S1. Supporting Information.

IMM-176-520-s002.pdf (76.4KB, pdf)

Lazana I., Simantirakis E., Kourous E., Daniil M., Ioannou D., and Vassilopoulos G., “Use of a Foamy‐Virus Vector System to Produce an ‘Off‐the‐Shelf’ Fcγ‐CR‐T Cell Product for the Treatment of Haematological and Solid Tumour Malignancies,” Immunology 176, no. 4 (2025): 520–529, 10.1111/imm.70018.

Funding: The authors received no specific funding for this work.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1. The binding capacity of various concentrations of the monoclonal antibodies (mAbs) Rituximab and Cetuximab to FV‐derived CD16‐CR T cells is illustrated. A dose‐dependent binding was noted, which starts to plateau at 0.1 μg/mL. This concentration (0.1 μg/mL) has been used in all subsequent experiments. μg/mL, micrograms per millilitre.

IMM-176-520-s001.docx (517KB, docx)

Data S1. Supporting Information.

IMM-176-520-s002.pdf (76.4KB, pdf)

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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