Preclinical evaluation of CAR20(NAP)-T cells for B cell lymphoma

doi:10.21203/rs.3.rs-4978161/v1

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Preclinical evaluation of CAR20(NAP)-T cells for B cell lymphoma

https://doi.org/10.21203/rs.3.rs-4978161/v1

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CD19-targeted CAR-T cell therapy has shown striking results against B cell malignancies, which has led to the approval of four CD19CAR-T cell products in the USA and EU. However, in long-term follow up evaluations it has become evident that many patients relapse after CD19CAR-T cell treatment and then in many cases present with CD19-negative tumors. In that case renewed CAR-T cell therapy targeting CD20 could be an option for lymphoma patients. Our previous study showed that CAR-T cells armed with immunomodulatory neutrophil-activating protein (NAP) from Helicobacter pylori (termed CAR(NAP)-T) can trigger the endogenous T-cell mediated immune response and further eliminate “CAR-target-antigen-negative” tumor cells. Here, we report the development of CD20-targeted (targeting moiety from Rituximab) CAR-T cells (CAR20-T cells), as well as the NAP-armed CAR20(NAP)-T cells and their pre-clinical evaluations in a murine lymphoma model. CAR20-T cells displayed efficient and specific cytotoxic potential against multiple human B cell lymphoma cell lines in vitro. In addition, primary mantle cell lymphoma cells isolated from a patient who relapsed after Rituximab treatment were also killed by CAR20-T cells. CAR20(NAP)-T cell treated mice showed delayed tumor growth and prolonged survival and NAP did not induce any severe toxicity. Human blood from healthy volunteers was exposed to recombinant NAP protein in an ex vivo human whole blood loop assay, without resulting in excessive cytokine release of immune cell activation, indicating a safe profile as a therapeutic transgene. CAR20(NAP)-T cells are worth further investigation especially in patients relapsing with CD19-negative tumors after CD19CAR-T cell therapy.

CD20

rituximab

chimeric antigen receptor

CAR-T cell

neutrophil-activating protein

tumor microenvironment

lymphoma

cytokine release

Cancer immunotherapy has demonstrated remarkable advances in the last decade. One of the key therapies that have contributed to this success is chimeric antigen receptor (CAR)-T cell therapy. CD19-targeting CAR-T cell therapy has been successfully implemented against therapy-resistant B cell malignancies with four products being approved in the USA and EU. The results have been especially striking when targeting acute lymphoblastic leukemia (ALL)[1–3], and diffuse large B cell lymphoma (DLBCL)[4–6]. Despite the encouraging outcomes of CAR T-cell therapy in both pivotal clinical trials and real-world scenarios, 30 to 60% of patients relapse after CAR T-cell therapy, and 10 to 20% of relapsed patients experience CD19-negative relapse[5, 7–10].

CD20 is expressed on B cells throughout most of their development and expression is retained in B-cell lymphomas. The CD20-targeting monoclonal antibody Rituximab was the first monoclonal antibody to be approved in oncology with excellent results for patients suffering from B cell malignancies[11, 12]. Rituximab together with CHOP (cyclophosphamide, hydroxydaunorubicin, oncovin and prednisone), the so-called R-CHOP regimen is the first-line treatment for several types of B-cell lymphomas[13–16]. At relapse, loss of CD20 or mutation of the Rituximab epitope is rarely the reason for failure of R-CHOP treatment[17]. Therefore, the Rituximab epitope of CD20 could still be targeted using a different therapeutic approach including CAR-T cells.

The Helicobacter pylori neutrophil-activating protein (NAP) is a major virulence factor of Helicobacter pylori. NAP promotes a T helper (Th) 1 polarized immune response, which is favorable in cancer treatment[18–20]. Also, NAP acts as a chemoattractant for innate immune cells, such as neutrophils and monocytes[21] and can both recruit and activate dendritic cells (DCs)[22]. We have shown that NAP, when released from NAP-armed CAR(NAP)-T cells, can be used as an immunomodulator to promote epitope spread and induce endogenous T cell responses towards CAR-target antigen-negative tumor cells [23].

The aim of the current study was to develop NAP-armed CD20-directed CAR-T cells which could be used to treat B cell lymphoma upon relapse from CD19CAR-T cell therapy. We show that CAR20-T cells displayed efficient and specific cytotoxicity in vitro against human lymphoma cell lines and patient-derived mantle cell lymphoma cells. Further, CAR20-T cells engineered to secret NAP (CAR20(NAP)-T cells) exhibit enhanced anti-tumor efficacy and can significantly prolong survival of mice with lymphoma. Arming CAR-T cells with NAP appears to be a safe strategy not causing excessive level of cytokine release in mice or over-activation of monocytes and granulocytes in human blood.

CAR20-T cells efficiently kill CD20-expressing lymphoma cells in vitro

Human CAR20-T cells including the single chain variable fragment (scFv) derived from Rituximab, and the intracellular activation domains from 4-1BB (CD137) and CD3ζ were produced from peripheral blood of healthy donors using a lentiviral vector also containing green fluorescent protein (GFP) to allow for detection of transduced T cells (Fig. 1A). T cells transduced with a lentiviral vector containing only GFP (termed Mock-T cells) was used as a negative control (Fig. 1A). Upon transduction, CAR20 was expressed on cell surface (Fig. 1B, C) and correlates nearly 100% with GFP expression (Fig. 1D). It has been shown that CARs, due to its intrinsic design, may induce antigen-independent tonic signaling and subsequent CAR-T cell exhaustion[24, 25]. Therefore we evaluated if CAR20-T cells displayed phenotypic exhaustion during in vitro culture. CAR20-T cells, as Mock-T cells, were mainly negative for activation marker (Fig. 1E), indicating that the design of the CD20-targeting CAR does not cause antigen-independent tonic signaling and CAR-T cell exhaustion.

To ivestigate if CAR20-T cells could specifically kill malignant B cell lines in vitro, CAR20-T cells were co-cultured with CD20⁺ (DB, U698, Raji, Daudi) and CD20⁻ (DG75) lymhoma cell lines were evaluated (Fig. 1F). CAR20-T cell killing was specific as all CD20-expressing lymphomas were killed, while the CD20⁻ lymphoma cell line DG75 was not (Fig. 1G, H; Suppl Fig. 1A, B). We also evaluated the cytotoxic capacity of CAR20-T cells in a clinically relevant setting, wherein CAR20-T cells were co-cultured with primary mantle cell lymphoma (MCL) cells isolated from a tumor biopsy of a patient that had become refractory to Rituximab treatment (Fig. 1I). Encouragingly, CAR20-T cells were able to efficiently kill CD20-expressing MCL cells as compared to Mock-T cells, evidenced as the significant reduction of CD20⁺ cells from the biopsy sample and release of IFN-γ (Fig. 1J-L).

In conclusion, CAR20-T cells could successfully be produced without induction of antigen-independent in vitro exhaustion. Further CAR20-T cells display potent cytotoxicity towards all tested CD20⁺ malignant B cell lines as well as primary CD20⁺ MCL cells in vitro.

Mouse mCAR20(NAP)-T cells exhibited enhanced anti-tumor efficacy in the experimental A20-hCD20 lymphoma model

To evaluate the therapeutic efficacy of CAR20-T cells in an immunocompetent mouse model, murine (m)CAR-T cells were engineered using retroviral vectors (RV) with the CAR20 functionally linked to the murine CD28 and CD3ζ intracellular signaling domains (Fig. 2A). We have previously shown that NAP-armed CAR-T cells improve therapeutic efficacy in several murine tumor models compared to conventional CAR-T cells[23]. To evaluate if NAP-armed mCAR20-T cells could improve therapeutic efficacy, a retrovirus vector for NAP-armed mCAR20-T cells was also constructed (Fig. 2A). As Rituximab only binds to human CD20, the murine B cell lymphoma A20 cell line was engineered to express human CD20, termed A20-hCD20, to allow evaluation of mCAR20-T cell treatment in vivo (Fig. 2B). First, the cytotoxic capacity of mCAR20-T cells was evaluated in vitro by co-culturing mCAR20-T, mCAR20(NAP)-T and Mock-T cells with A20-hCD20 tumor cells at different effectors to target cell ratios. Both mCAR20-T and mCAR20(NAP)-T cells could efficiently kill the A20-hCD20 target cells (Fig. 2C) and secreted IFN-γ (Fig. 2D) upon antigen encounter. There was no apparent difference between mCAR20-T and mCAR20(NAP)-T cells in terms of cytotoxicity and IFN-γ, indicating that insertion of NAP did not negatively affect the in vitro antitumor efficacy, which is in accordance with our previous findings[23]. Further, in a xenograft setting, systemically injected human CAR20(NAP)-T cell treatment slowed down human tumor growth, prolonged the survival and cured some mice, supporting in vivo cytotoxicity of CAR20(NAP)-T cells (Suppl Fig. 2A-C).

To be able to fully evaluate the potency of NAP-armed CAR20-T cells in vivo, we investigated the therapeutic efficacy of CAR-engineered T cells in an immunocompetent mouse model with subcutaneous A20-hCD20 lymphoma. Mice were treated intravenously (i.v.) with mCAR20, mCAR20(NAP) or Mock-T cells 10-, 14-, and 18-days post tumor implantation (Fig. 2E). No visible toxicity was observed after treatment and the mice maintained a stable weight during treatment (Fig. 2F). The presence of human CD20 in A20-hCD20 lymphomas rendered them immunogenic as nearly half of the mice became tumor-free even in the Mock-T treated group. However, mCAR20-T and mCAR20(NAP)-T treatment still improved the therapeutic outcome as the average tumor growth was significantly delayed in comparison to mice treated with Mock-T (Fig. 2G). Incorporation of NAP into mCAR20-T cells further delayed tumor growth (Fig. 2G) and significantly prolonged the survival of mice compared the Mock-T cell treated mice (Fig. 2H). In addition, there were more tumor free mice after both mCAR20-T and mCAR20(NAP)-T treatment in comparison to Mock-T cell treated mice (Fig. 2I-K).

Treatment-related toxicity was not observed in CAR20(NAP)-T treated mice

There were no detectable side effects in mice with A20-hCD20 tumors upon treatment with CAR20(NAP)-T treatment, observed as stable body weight after treatment (Fig. 2F). However, as CAR20(NAP)-T cells can only recognize human CD20 and cannot be activated by normal mouse B cells and NAP expression is induced upon antigen recognition, it is difficult to assess potential side effects from a potentially broader systemic secretion of NAP in the A20-hCD20 model. To evaluate potential side effects of NAP we therefore treated A20 (CD19⁺) lymphoma-bearing mice with mCAR19-T and mCAR19(NAP)-T cells, targeting murine CD19 and thus also normal mouse B cells (Fig. 3A). After treatment, the heart, kidney, liver, lung, brain, and spleen were collected to evaluate potential tissue damage (Fig. 3A). No visible symptoms were observed in mice treated with either of CAR-T or CAR(NAP)-T cells and the body weight of these mice remained stable throughout the study (Fig. 3B). There were no signs of lesions in kidney, liver, lung, brain, and spleen after treatment. In the heart there were mild lesions that might suggest myxomatous degeneration of cardiac valves (Fig. 3C-K), observed in all treatment groups, including Mock-T cell treated animals. Thus, it is unlikely that the lesions were associated with systemic release of NAP. In conclusion, arming the CAR-T cells with NAP did not change the safety profile of CAR-T cell treatment.

Recombinant NAP protein does not cause toxicity in human blood

In a clinical setting, NAP would be secreted systemically if CAR20(NAP)-T cells encounter healthy B cells in the blood of patients. We therefore sought to investigate the potential toxicity due to systemic release of NAP. Recombinant NAP protein was incubated into a whole human blood loop system[26] to investigate if and to which extent NAP induces cytokine release, complement activation and immune cell activation in fresh circulating blood from healthy human donors (Fig. 4A). Two concentrations of NAP proteins were evaluated in the blood loop system, 5 and 25µg/ml. Alemtuzumab (anti-CD52, 3µg/ml) was used as reference control since it is used for treatment of B cell chronic lymphocytic leukemia with manageable adverse events also with respect to cytokine release [27, 28]. Lipopolysaccharide (LPS) was used as a positive activating control as it is one of the most potent innate immune activating stimuli that triggers strong release of TNFα and IL-6 from innate cells such as monocytes[29].

We observed an increased level of IL-6 over PBS control with an average of 3.9-fold increase at 5µg/ml of NAP and 4.3-fold increase (p < 0.05) at 25µg/ml of NAP (Fig. 4B). The levels of IL-8 and TNFα were increased over PBS control at both NAP concentrations, with an average of 1.7-fold and 4.3-fold increase of IL-8 (p < 0.05 at 5µg/ml of NAP and p < 0.001 at 25µg/ml of NAP) and an average of 2.3-fold and 4.4-fold increase of TNFα (p < 0.001 at 25µg/ml of NAP) (Fig. 4C, D). Neither concentration of NAP induced quantifiable level of IFN-γ and IL-2 in human blood, as all values were below the lower limit of quantification (LLOQ) (Fig. 4E, F). In all cases, NAP induced less cytokine release than the reference Alemtuzumab (Fig. 4B-F). Additionally, we observed a modest increase of complement split product C3a for NAP over PBS control (Fig. 4G), while the split product C5a was significantly increased at both NAP concentrations (p < 0.05) (Fig. 4H). This indicates that NAP, at the tested concentration, cannot activate the complement system to a functional level as compared to Alemtuzumab.

Next, we analyzed the ability of recombinant NAP protein to induce the activation of immune cells. Platelet activation was assessed based on the level of CD62P on CD41⁺CD45⁻ cells (Fig. 4I). The expression level of CD69 and CD107a were used to access the activation of T cells (CD3⁺CD56⁻), B cells (CD3⁻CD19⁺) and NK cells (CD3⁻CD56⁺). The expression of CD83 and CD11b were used to assess the activation of monocytes (CD14⁺CD66b⁻) and granulocytes (CD14⁻CD66b⁺). In short, we did not see any significant NAP-mediated activation of platelets, T cells, B cells, NK cells, monocytes or granulocytes at either of the two assessed concentration levels (Fig. 4J-N). Alemtuzumab activated T cells, NK cells and granulocytes (Fig. 4J, K, M). None of the treatments had any effect on blood cell counts (Suppl Fig. 3A-H). In conclusion, NAP induced concentration-dependent cytokine release, and a slight complement activation. The activation levels were lower than the reference control Alemtuzumab, which side-effects is manageable in the clinic.

CD19CAR-T cell therapy has shown successful results against relapsed/refractory (r/r) B cell malignancies[4-6]. However, follow up has shown that 10-20%patients acquire secondary resistance which is often associated with downregulation or loss of CD19[5, 9]. CD20 has been proposed as an alternative CAR-target for treatment of large B cell lymphoma (LBCL) as it remains expressed after CD19CAR-T cell therapy[9]. In addition, CD20 alterations or loss are rare in LBCL despite the usage of Rituximab anti-CD20 antibody therapy in standard of care[9, 17].

The single-chain variable fragment derived from the anti-CD20 antibody Rituximab was used to construct CAR20-T cells. CAR20-T cells displayed efficient and specific cytotoxic potential against multiple CD20-expressing lymphoma cells in vitro. In the clinic, Rituximab together with chemotherapy (R-CHOP) is used to treat patients with LBCL. CD20-targeting antibody therapy rarely alters CD20 expression[9, 17]. In line with this, our CAR20-T cells displayed cytotoxicity against primary mantle cell lymphoma cells from a patient relapsed after R-CHOP despite targeting the Rituximab epitope. Even though the tumor-cell killing is performed ex vivo, this result indicated clinical feasibility for our CAR20-T cells.

Selection pressure from CAR20-T cell therapy would most likely lead to modification or loss of CD20 as commonly seen for CD19 after CD19CAR-T cell therapy[5, 9]. To overcome antigen-loss, CAR-T cells simultaneously targeting multiple antigens have been developed[30-32]. However, dual antigen-loss has been observed[32, 33] indicating the need for new approaches to prevent relapse due to antigen loss. We have previously shown that arming CAR-T cells with inducible secretion of the bacterial-derived immune-stimulating factor NAP can induce bystander immunity and epitope spread and thereby elimination of CAR-target-negative tumor cells[23]. In this study, mCAR(NAP)-T cells could slow down tumor growth and prolong survival in lymphoma bearing mice without inducing toxicity . Improvement of mCAR20-T cell therapy upon incorporation of NAP implies that the therapy could be evaluated for patients with CD20-positive relapsed r/r B cell lymphomas, possibly after relapse from CD19CAR-T cell treatment.

No visible symptoms or weight loss were observed after mCAR20- or mCAR20(NAP)-T cell therapy indicating a safe profile of mCAR20(NAP)-T cell treatment (Figure 2F). However, Rituximab only recognizes human CD20 and thus potential toxicity of systemic NAP, upon encounter of healthy B cells in the circulation, could not be evaluated in this murine model. Therefore, to assess safety of NAP-armed CAR-T cells, murine CD19-targeting CAR19-T cells were evaluated in the murine A20 lymphoma model. Only the heart displayed mild lesions suggesting potential myxomatous degeneration of the cardiac valves (Figure 3). It has been reported in some studies that CAR-T cell therapy can affect the cardiovascular system[5, 34]. Importantly in our study, potential cardiac valve effects were observed in all treatment groups indicating the toxicity phenomenon is most likely not CAR-T cell treatment related.

We further investigated the possibility of NAP-induced systemic toxicity in human blood using the blood loop assay, which includes a specifically designed cytokine panel, can be applied in clinical settings to predict adverse events like cytokine release syndrome (CRS).[35]. A panel of cytokines, complement activation markers and cell activation markers were analyzed. NAP did not induce lymphocyte activation as compared to Alemtuzumab (Figure 4I-K). NAP increased the plasma levels of IL-6, IL-8 and TNFα, which have all been implicated in CRS in a concentration-dependent manner (Figure 4A-C). Alemtuzumab, which is approved for the treatment of chronic lymphocytic leukemia and multiple sclerosis and frequently induces CRS which can be managed by corticosteroid treatment[28, 36]. As NAP induces lower levels of cytokine release compared to Alemtuzumab (Figure4A-E), we predict that cytokine release induced by NAP, secreted from CAR-T cells, should be manageable in the clinic.

In summary, we have developed efficient CD20-targeting CAR-T cells that once armed with NAP display improved therapeutic potential in vivo without altering the safety profile. NAP further displayed a safe profile in a human blood loop assay. This study paves the way for further evaluation of the CAR20(NAP)-T in patients with refractory and relapsed B-cell lymphoma.

Cell lines and primary cells culturing conditions

Human lymphoma cell lines DB, and DG-75[37] (Kind gift from Dr G Klein, Karolinska Institute, Stockholm, Sweden), U-698[38] (Kind gift Kenneth Nilsson, Uppsala University), Daudi (ATCC CCL-213), Raji (Sigma Aldrich) were cultured in RPMI 1640 supplemented with 10% heat-inactivated fetal bovine serum (FBS), 1% penicillin/streptomycin (PeSt) and 1% sodium pyruvate. All reagents were from Invitrogen. All lymphoma cell lines were modified to stably express green fluorescent protein, firefly luciferase (FLuc) and puromycin (for selection) by lentiviral transduction (Magnus Essand, Addgene, Plasmid #80389)[39]. Cells expressing transgenes were selected using 5µg/ml puromycin.

Mouse B-cell lymphoma cell line A20 (ATCC TIB-208) was cultured in culture medium (RPMI 1640) containing 10% heat-inactivated FBS, 1% (PeSt) and 1% sodium pyruvate. To generate A20 cells stably expressing human CD20 (termed as A20-hCD20), A20 cells were transduced with a lentiviral vector encoding hCD20, FLuc and puromycin separated by a self-cleaving peptide (T2A and F2A). A20-hCD20 cells were selected using 3µg/ml puromycin.

Peripheral blood mononuclear cells (PBMCs) were isolated by Ficoll-Paque (GE Healthcare Life Science) from fresh buffy coats taken from healthy donors, collected from the Blood Center at Uppsala University Hospital. They were used as a source for human T cells, which were cultured in RPMI 1640 supplemented with 10% FBS, 1% PeSt and 1% sodium pyruvate with 50IU/ml IL-2 (Proleukin, Novartis) if not stated otherwise.

Retro- and lentiviral vector construction and production

To create lentiviral vectors encoding CAR20 the variable region sequence from Rituximab was incorporated into a CAR cassette with the signaling domain of CD3zeta and the co-stimulatory signaling domain of CD137 as previously described[40]. The CAR constructs were cloned into a third-generation self-inactivating (SIN) lentiviral vector (SBI, System Biosciences) under the control of elongation factor-1 alpha (EF1a) promoter. Green fluorescence protein (GFP) was inserted after the CAR construct using a T2A self-cleaving peptide in both plasmids to allow detection of CAR-T cells. A mock construct only encoding GFP under the control of the EF1a promoter was used as a control. All sequences were purchased from GenScript. Production of third generation viral particles has been previously described[41]. The retroviral vector encoding human CAR20 sequence was purchased from BioNTech and used for therapeutic efficacy studies in xenograft Daudi model.

The variable region of Rituximab was incorporated into a murine CAR construct containing murine CD3ζ and murine CD28 signaling domains, has been described previously[42]. To construct retroviral vector RV(mCAR20), the above constructs were subcloned into mouse stem cell virus-based retroviral vector, pMIG-W (Luk Parijs, Addgene, plasmid #12282)[43]. The RV(mCAR19) and RV(mCAR19-NAP) constructs have been described previously[23]. An empty retroviral vector was used as Mock control. The NAP sequence was inserted in the orientation opposite the CAR20 cassette and under the control of murine NFAT-IL-2 promoter to construct retroviral vector RV(mCAR20-NAP). Retrovirus was produced using the packaging plasmid pCL-Eco in the Gryphon-Eco retroviral packaging cell line (Allele Biotechnology).

Human T cell engineering, enrichment and expansion

Human PBMCs, see above, were activated using OKT-3 (100ng/ml, BioLegend San Diego, CA) or TransAct (1:100, Miltenyi Biotec) and IL-2 (100 IU/ml, Proleukin) for 3 days at a concentration of 2x10⁶ cells per ml. Following activated T cells (1x10⁶ cells) were re-suspended in 20µl concentrated lentivirus together with 10mg/ml protamine sulfate (Sigma-Aldrich) and IL-2 (100 IU) and incubated for 4hrs at 37°C. The following day T cells were transduced in the same manner. After transduction T cells were cultured (in culture medium supplemented with 50IU/ml IL-2) for 7 days before CAR-T cells were enriched by sorting out GFP⁺ cells (BD FACSAriaIII, BD Biosciences). Sorted cells were expanded using rapid expansion protocol (REP) previously described[44].

Mouse T cell activation and transduction

Spleens were collected from BALB/c mice (Janvier) and mashed against a 70µm cell strainer (Coring) to create single cell suspension. After resuspended in red blood cell lysis buffer (ACK lysing buffer, Invitrogen) for 5min, splenocytes were activated using Concanavalin A (2µg/ml) (Sigma Aldrich) and murine IL-7 (Miltenyi Biotec) in RPMI 1640 containing 10% FBS, 1% PeSt and 1% sodium pyruvate. Two days post activation splenocytes were transduced with retrovirus as previously described[23].

T cell cytotoxicity assay and IFN-γ ELISA

Human T cell cytotoxicity assay:

The cytotoxicity assay was performed using luciferase-expressing target cells. CAR-engineered human T cells were rested for 3d after rapid expansion in a medium containing a low dose of IL-2 (20 IU/ml) before performing the cytotoxicity assay. The T cells were co-cultured with luciferase-tagged target cells (DG75, DB, U698, Raji and Daudi, at 3:1 ratio) in a total volume of 200μl, in a round-bottom 96-well plate. To detect the luciferase expression and activity, ONE-Glo luciferase assay system (Promega) was added according to manufacturer’s protocol. Specific lysis of target cells was calculated by comparing luminescence of co-cultured samples to untreated target cells.

Murine T cell cytotoxicity assay:

The cytotoxicity assay was performed using luciferase-expressing target cells. Murine splenocytes were harvested, activated and transduced with retroviral vectors as above described. The engineered-T cells were co-cultured with firefly luciferase-tagged A20-hCD20 target cells at different ratios (50:1, 20:1, 5:1 and 1:1) in a round bottom 96-well plate (total volume 200µl). After 5 days of co-culture, 80µl supernatant was collected for IFN-γ analysis determined by an ELISA kit (Mabtech). Luciferase expression and activity (as an indicator of target cell viability) were determined by using ONE-Glo luciferase assay system (Promega) according to manufacturer’s protocol. Specific lysis of target cells was calculated by relating luminescence of co-culture with target cells alone.

T-cell cytotoxicity assay towards patient samples

Biopsy samples containing primary mantle cell lymphoma (MCL) cells were either left untreated or co-cultured together with engineered-T cells at a 1:1 (1=25 000 cells) ratio for 3 days in round bottom 96-well plate in a total volume of 200µl per well. After 2 days co-culture, supernatant was collected for analysis of IFNγ secretion by an ELISA kit (Mabtech). After 3 days of co-culture, the remaining amount of CD20⁺ cells (as an indicator of target cell viability) was assessed by flow cytometry. Specific lysis of target cells was shown as the proportion of CD20⁺ cells normalized to the one at the start of the experiment.

Cytotoxicity assay using Incucyte

Measurement of immune cell killing of target tumor cells was analyzed with the Incucyte® Zoom image analysis software (Essen BioScience). Briefly, one image per well in a 24-hour repeat scanning was schedule with the Phase contrast, green and red channels in 10x objective. Images were taken every 30 minutes with the standard scan schedule and “Basic Analyzer” module. Image collections and processing definitions for every cell line and assay were stablished following the suggested procedures by the manufacturer. Processing definitions were stablished with phase and fluorescence masks for scarlet expression to quantify target cell proliferation, determined as total red fluorescence confluence. Target cell killing by CAR-T cells was determined by comparing red fluorescent signal of co-cultured (at the effector to target ratio 5:1) samples to untreated target cells and reported as relative cell viability.

Flow cytometry

In order to assess transduction efficiency, CAR and GFP expression of engineered-T cells were detected by flow cytometry. The T cells were stained with CD3-BB700 (Clone SP34-2) and anti-mouse IgG (H+L)-AF647 (to detect CAR expression, A21237, Thermo Fisher Scientific).

Human CD20 expression in human cell lines and in the modified A20-hCD20 cells was assessed by staining the cells with anti-human CD20-APC (Clone 2H7), isotype was used as control (Mouse IgG2b,κ). Stained cells were analyzed by flow cytometry using BD FACSCanto II (BD Biosciences) or CytoFLEX LX (Beckman Coulter) and analyzed using FlowJo_v10.8.1 (FlowJo LLC).

Animal experiment

Survival experiment

Daudi model: NOD-SCID mice (8 weeks) were implanted intravenously (i.v.) with 1×10⁶ firefly luciferase (fLuc) expressing Daudi cells. Mice were then treated with two doses (2×10⁶ cells/dose) intravenously on 4 and 8 days post tumor implantation. The tumor progression was monitored by in vivo imaging system (IVIS) Lumina III (PerkinElmer). Mice were intraperitoneal (i.p.) injected with 100 μL of 30 mg/mL of D-Luciferin potassium salt (Perkin Elmer) and imaged 10min after substrate injection. Mice were monitored regularly for their health status and euthanized when reaching the humane endpoint.

A20-hCD20 model: Female BALB/c mice (8 weeks) were implanted subcutaneously (s.c.) with 2×10⁵ murine lymphoma A20 expressing human CD20 (A20-hCD20). The mice were treated with 3×10⁶ T cells intravenously (i.v.) 10-, 14-, and 18-days post tumor implantation. The mice were monitored regularly and euthanized when reaching the humane endpoint. Mice were sacrificed when reaching the humane endpoint (a tumor size reach 1cm³). The experiment was performed twice, and all data were pooled together (Mock-T: n=20, mCAR20-T: n=20, mCAR20(NAP)-T: n=19).

The mice were monitored for tumor growth until reaching the humane endpoint volume 1000 mm³. The tumor size was calculated as: volume=length×width²×π/6. The time to endpoint (TTE) for each mouse was calculated as TTE= [log (EPV)-b]/m. The constant b is the intercept and m are the slope of the line obtained by linear regression (time vs tumor volume) of a log-transformed tumor volume data set, which comprised of the first measured tumor volume when EPV was exceeded and three consecutive tumor volumes immediately prior to the attainment of EPV. Any animal that died from treatment-related causes was assigned a TTE value equal to the day of death. Any animal that died from non-treatment-related causes was excluded from the analysis. Survival curve was generated based on the TTE value using the Kaplan-Meier method and compared using the log-rank (Mantel-Cox) test.

Safety assessment

Female Balb/c mice (3 mice/group) were subcutaneously implanted on day 0 with A20 tumor (murine lymphoma cell line A20) and treated intravenously with Mock-T, mCAR-T or mCAR(NAP)-T cells (targeting murine CD19) on day 10, 14 and 18. Different organs (brain, heart, kidney, lung, liver, and spleen) were collected on day 20 and, fixed in 4% formalin, embedded paraffin and sent to BioVet AB for pathologic evaluation. Sections were 3-4 um thin and stained with Heamatoxylin-Eosin (HE). Photomicrographs were taken when the observed lesions appeared.

Whole blood loop analyses

Fresh whole blood was taken from ten healthy volunteers (D1-D10) and immediately mixed with a low amount of soluble heparin. Then whole blood was immediately transferred to heparin pre-coated plastic tubes forming the loops, followed by administration of the test items, 5μg/ml NAP proteins and 25μg/ml NAP proteins. PBS was used as negative control and 3μg/ml Alemtuzumab (Lemtrada, Sanofi) was used as positive antibody, and 3μg/ml LPS (Sigma-Aldrich, Sweden) was used as positive control. Additionally, fresh blood from each donor was used for hematology measurements directly after blood collection (described as pre-loop baseline). Additionally, blood collected directly from donors was processed to plasma and included in cytokine and complement analysis. Blood samples were collected at 15 minutes for complement analysis and immune cells activation, while blood samples collected at 4 hours were analyzed for hematology analysis, immune cell activation and cytokine analysis.

For hematology analysis, it included the parameters: red blood cell (10¹²/L), platelets (10⁹/L), white blood cell (10⁹/L), differential neutrophil (10⁹/L), lymphocyte (10⁹/L), monocyte (10⁹/L), eosinophil (10⁹/L) and basophil (10⁹/L). These parameters were measured at 0h (pre-loop baseline) and 4h time-point by the Hematology Analyzer Sysmex XN-L350 (Sysmex). Hematology analysis of zero time point samples was performed as three subsequent measurements, while samples collected in the loop system at 4 hours were done as single measurements.

For cytokine and complement analysis, plasma harvested at 0 (pre-loop baseline) and 4-hour time-point was analyzed for IFN-γ, TNF-α, IL-2, IL-6 and IL-8 with a Mesoscale V-plex kit (Meso Scale Discovery) according to the manufacturer's instructions. Complement analysis (C3a and C5a) was performed on plasma collected at 0h (pre-loop baseline) and 15min time-point with ELISA kits (Raybiotech) according to the manufacturer’s instructions. Detection limit or limit of detection (LOD) is the smallest concentration of an analyte from which it is possible to deduce the presence of an analyte in the test sample, while quantification limit or limit of quantification (LOQ) is the smallest concentration of an analyte that can be determined with the specified degree of accuracy and precision. Lower limit of detection (LLOD) was calculated by MSD software and defined as 2.5xSD above the zero calibrator (Standard-8).

For immune cells activation, blood was collected at the 0, 15min and/or 4h time points, and the activation of immune cells were analyzed by flow cytometry (CytoFLEX LX, Beckman Coulter). The platelets were gated as CD41⁺ CD45^- cells and analyzed the activation marker CD62P. The lymphocytes were first identified in the forward versus side scatter (FSC/SSC) plot and further gated as T cells (CD3⁺CD56^-), NK cells (CD3^-CD56⁺) and B cells (CD3^-CD19⁺). The activation phenotype of lymphocytes was determined by staining by CD69 and CD107a. Granulocytes and monocytes were as well identified in FSC/SSC plot and further defined as granulocytes (CD14^-CD66b⁺) and monocytes (CD14+CD66b^-), respectively. The activation markers were analyzed by staining with CD11b and CD83.

Ethical approval

The Uppsala Research Animal Ethics Committee has approved all the animal studies (5.8.18-19434-2019). The human buffy coat obtained from healthy donors were anonymized. The patient-derived mantle cell lymphoma cells were collected under ethical permit: EPN 233-2014.

Statistical analysis

Two-way ANOVA with Tukey multiple comparisons test was used to compare means between more than two experimental groups. For statistical analysis between two experimental groups, a non-parametric unpaired t-test was used to compare means between experimental groups. Statistical significance of survival (Kaplan-Meier) a log-rank test was used. A p value of less than 0.05 was considered as statistically significant.

Author Contributions

Conception and Design: JM, TS, HSK, GE, DY, ME; Acquisition of Data: JM, TS; CJ Analysis and Interpretation of Data: JM, TS; Writing, Review, and/or Revision of the Manuscript: JM, TS, CJ, PD, DY, ME

Conflicts of interest

The authors declare no financial conflict of interest

Acknowledgements

The authors wish to thank Minttu-Maria Martikainen (Uppsala University) for assistance in animal work and the staff at BioVis for flow sorting. The authors also want to express their sincere gratitude to Angelica Loskog and Tanja Lövgren for valuable discussions and input to the project. The authors would like to thank Ingrid Thörn for the possibility to evaluate the newly developed CAR-T cells against a primary mantle cell lymphoma sample. Funding: This work was supported by research grants from the Swedish Research Council: 2019-01326 (ME), the Swedish Cancer Society: 19-0184Pj (ME), 19-0188Us (ME), 22-2241-Pj (ME), 22-2229Pj (DY), the Swedish Childhood Cancer Fund: PR2020-0167 (ME), PR2022-0105 (DY) and VINNOVA 2021-04488 (DY).

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No competing interests reported.

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Preclinical evaluation of CAR20(NAP)-T cells for B cell lymphoma

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Abstract

Figures

Introduction

Results

Mouse mCAR20(NAP)-T cells exhibited enhanced anti-tumor efficacy in the experimental A20-hCD20 lymphoma model

Treatment-related toxicity was not observed in CAR20(NAP)-T treated mice

Recombinant NAP protein does not cause toxicity in human blood

Discussion

Materials and Methods

Declarations

References

Additional Declarations

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