- Email: [email protected]
Blocking CD38-driven fratricide among T cells enables effective antitumor activity by CD38-specific chimeric antigen receptor T cells
Journal of Genetics and Genomics 46 (2019) 367e377
Contents lists available at ScienceDirect
Journal of Genetics and Genomics Journal homepage: www.journals.elsevier.com/journal-of-geneticsand-genomics/
Original research
Blocking CD38-driven fratricide among T cells enables effective antitumor activity by CD38-specific chimeric antigen receptor T cells Zhitao Gao, Chuan Tong, Yao Wang, Deyun Chen, Zhiqiang Wu*, Weidong Han* Department of Molecular Biology, Institute of Basic Medicine, School of Life Sciences, Medical School of Chinese PLA, Chinese PLA General Hospital, Beijing, 100086, China
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 April 2019 Received in revised form 9 June 2019 Accepted 17 June 2019 Available online 13 August 2019
Chimeric antigen receptor T-cell (CAR T) therapy is a kind of effective cancer immunotherapy. However, designing CARs remains a challenge because many targetable antigens are shared by T cells and tumor cells. This shared expression of antigens can cause CAR T cell fratricide. CD38-targeting approaches (e.g., daratumumab) have been used in clinical therapy and have shown promising results. CD38 is a kind of surface glycoprotein present in a variety of cells, such as T lymphocytes and tumor cells. It was previously reported that CD38-based CAR T cells may undergo apoptosis or T cell-mediated killing (fratricide) during cell manufacturing. In this study, a CAR containing a sequence targeting human CD38 was designed to be functional. To avoid fratricide driven by CD38 and ensure the production of CAR T cells, two distinct strategies based on antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) were used to block CD38 or the CAR single-chain variable fragment (scFv) domain, respectively, on the T cell surface. The results indicated that the antibodies or proteins, especially the antibody MM27, could affect CAR T cells by inhibiting fratricide while promoting expansion and enrichment. Anti-CD38 CAR T cells exhibited robust and specific cytotoxicity to CD38þ cell lines and tumor cells. Furthermore, the levels of the proinflammatory factors TNF-a, IFN-g and IL-2 were significantly upregulated in the supernatants of A549CD38þ cells. Finally, significant control of disease progression was demonstrated in xenograft mouse models. In conclusion, these findings will help to further enhance the expansion, persistence and function of anti-CD38 CAR T cells in subsequent clinical trials. Copyright © 2019, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, and Genetics Society of China. Published by Elsevier Limited and Science Press. All rights reserved.
Keywords: Chimeric antigen receptor CD38 T cells Immunotherapy CD38 antibody
1. Introduction Adoptive cell therapy (ACT) with genetically modified T cells expressing a chimeric antigen receptor (CAR), which are artificially engineered receptors expressed on the cell surface of T cells with non-HLA-restricted tumor antigens, allows CARs to activate T cells and specifically guide them to tumor cells to perform their function (Han et al., 2018; Zhang and Kasi, 2018). However, designing CARs remains a challenge because of the shared expression of many targetable antigens between normal cells and tumor cells, including targets expressed on the T cell surface, such as NKG2D, CS1, CD7, and CD5. This shared expression of antigens can cause fratricide of CAR T cells, inhibiting their proliferation and viability
* Corresponding authors. E-mail addresses: [email protected] (Z. Wu), [email protected] (W. Han).
and causing unpredictable on-target, off-tumor toxicity in clinical studies (Brudno and Kochenderfer, 2019). Human CD38 antigen (45 kDa) is a type II transmembrane glycoprotein that is expressed by monocytes, plasma cells and immature cells and activates T and B lymphocytes (Atanackovic et al., 2016; Bonello et al., 2018; Morandi et al., 2018). In the normal hematopoietic system, CD38 is expressed only by precursors that are “lineage committed” (Mihara et al., 2012) and is rarely expressed by multipotent stem cells and nonhematopoietic tissues. However, it is widely expressed on various types of cancer cells. CD38 is overexpressed in several lymphocyte malignancies, including B-cell non-Hodgkin lymphoma (B-NHL) (Mihara et al., 2010), multiple myeloma (MM), and collagen disease. CD38 is also expressed in some refractory and/or relapsed acute lymphoblastic leukemias (ALL) (Blatt et al., 2018; Naik et al., 2018), chronic lymphocytic leukemia (CLL) (Mele et al., 2018) and non-Hodgkin lymphoma (NHL) associated with CD38-positive cells (Zaja et al., 2017), which are especially associated with relapse and/or
https://doi.org/10.1016/j.jgg.2019.06.007 1673-8527/Copyright © 2019, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, and Genetics Society of China. Published by Elsevier Limited and Science Press. All rights reserved.
368
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
refractory cancer after anti-CD19 CAR T cell therapy. Furthermore, sustained expression of CD38 on tumor cells is associated with a poor prognosis (Pick et al., 2018). On the basis of previous studies, CD38 is regarded as an attractive target, and targeting CD38 is a potential treatment strategy for hematological malignancies. An anti-CD38 antibody (daratumumab) has been approved for the treatment of MM by the Food and Drug Administration (FDA) (Chehab et al., 2018; van de Donk and Usmani, 2018), and preclinical and clinical studies have demonstrated the efficacy and safety of the anti-CD38 antibody (Plesner and Krejcik, 2018; van de Donk, 2018). CD38 plays an important role in CAR T cell immunotherapy as a molecular target. However, CD38 is also present on the surface of NK cells and T cells. Thus, these cells may undergo suicide upon the transduction of CAR T cells. In this study, a lentiviral vector carrying a CAR comprising an anti-CD38 single-chain variable fragment (scFv) domains was designed and the antitumor activity of antiCD38 CAR-modified T cells was evaluated. As previously reported, activated T cells can also express CD38, suggesting that these T cells may undergo apoptosis or T cell-mediated killing (fratricide) during cell manufacturing (McHayleh et al., 2019; Roddie et al., 2019). To inhibit target-driven fratricide, two different strategies based on CD38 proteins or blocking antibodies were investigated. It was found that antibodies or proteins can affect anti-CD38 CAR T cells by inhibiting fratricide while promoting expansion and enrichment. Further, human T cells have been demonstrated to be resistant to CD38-positive cells with high efficiency both in vivo and in vitro in the present of anti-CD38 CAR T cells. These strategies have been incorporated into on-going clinical trials (NCT03754764) testing the efficacy of this CAR-modified T-cell immunotherapy for the treatment of relapsed and/or refractory leukemia associated with CD38-positive cells, especially for the treatment of relapsed and/or refractory disease after CAR T cell therapy.
2. Results 2.1. An anti-CD38 CAR design and the expression of the CAR on the surface of T cells To test whether normal T cells can recognize tumor cells via a CD38-specific CAR, we created a CAR construct with the human anti-CD38 scFv linked to a CD8a hinge and transmembrane region followed by the human 4-1BB (CD137) and CD3z intracellular signaling motifs, as shown in Fig. 1A. Similarly, MOCK T cells were constructed and used as a control. HEK-293T cells cotransfected with packaging plasmids were used to prepare lentiviral particles encoding the CAR plasmid (Feng et al., 2017). To confirm anti-CD38 CAR expression in transduced T cells, a CD38 Fc-tagged protein was used, followed by incubation with a goat anti-Fc antibody conjugated with APC that cross-reacts with the anti-CD38 CAR. The expression of anti-CD38 CAR was then analyzed by flow cytometry. The anti-CD38 CAR T cell transduction efficiency reached 90.53% ± 8.72% after 15 days (Fig. 1B and C). To assess cytotoxicity, we cocultured anti-CD38 CAR T cells with CD38-positive RPMI 8226 cells (Fig. 1D). After 24 h, 62.40% ± 6.73% of the RPMI 8226 cells were lysed by the anti-CD38 CAR T cells (Fig. 1E and F) at an effector-to-target (E:T) ratio of 5:1, while little lysis was observed in the coculture with MOCK T cells. In contrast, no cytotoxicity was detected in a K562 cell assay, as K562 cells lack CD38 expression. In an assay with CD38-positive primary tumor cells (Fig. 1G), anti-CD38 CAR T cells were incubated with the primary cells at an E:T ratio of 1:1 for 6e48 h. The efficiency of tumor cell lysis by the CAR T cells reached 22.76% at 24 h and 40.44% at 48 h (Fig. 1H).
2.2. CD38-driven fratricide of anti-CD38 CAR T cells To confirm the occurrence of cytotoxicity via CAR T cell fratricide, flow cytometry was carried out to detect the expression of CD107a during anti-CD38 CAR T cell expansion over 6 days (Fig. 2A). Normally, CD107a is regarded as a marker of CAR T cell degranulation. Once CAR T cells contact their target antigen, they are rapidly activated, express CD107a, and undergo degranulation (Kloss et al., 2019). However, human CD38 antigen (Ag) was prevalently expressed on T lymphocytes and caused CAR T cell activation. Cytotoxicity increased with increasing expression of CD107a, which may be associated with the loss of the CD38 Ag from the cell surface (Fig. 2B), and many dead cells and much debris were simultaneously observed (Fig. 2C). The decrease in CD38 expression on CAR T cells or the lysis of CD38þ CAR T cells by activated anti-CD38 CAR T cells appears to lead to a “CAR T cell - autophagic and autostimulatory/enrichment” cycle. In initial experiments, the population of anti-CD38 CAR T cells was measured during the cell expansion process. We found that compared with MOCK T cells, anti-CD38 CAR T cells could not achieve a rapid expansion ratio. Although T cells expressing this CAR have been successfully generated, there were significant differences in enrichment and proliferation between the CAR T cell group and the MOCK T cell group (Fig. 2D). We hypothesized that target-driven fratricide caused the loss of cell viability among the anti-CD38 CAR T cells. 2.3. Antibodies or proteins promote the enrichment and expansion of CAR T cells To avoid fratricide driven by targeting and to improve anti-CD38 CAR T cell production for clinical application, two distinct methods were investigated. The first approach focused on CD38 proteins to block the anti-CD38 CAR scFv. The second strategy involved antibody blockade of the CD38 Ag during CAR T cell production. Therefore, we tested whether an antibody or a protein could abrogate anti-CD38 CAR T cell-mediated fratricide during T cell culture. Antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) were added during the culture of anti-CD38 CAR T cells, and MOCK T cell and no additive CAR T cell groups were established at the same time. After 15 days of cell culture and expansion, the anti-CD38 CAR T cell transduction efficiency ranged from 58.61% to 94%, and the mean MOCK T cell transduction efficiency reached 83.57% ± 6.57%. In particular, the total cell number of the MM27 group was highly enriched ((102.36 ± 10.98) 106), and the transduction efficiency reached 76.49% ± 4.83% (Fig. 3A and B). Similarly, the total number of MM12T group reached (87.64 ± 5.79) 106, and the transduction efficiency reached 67.77% ± 5.16%. Furthermore, compared with the no additive CAR T cell group, the two groups treated with a protein (H02H or H08H) to block the CAR scFv domain also exhibited significantly improved CAR T cell expansion and transduction efficiency. Since the antibodies used blocked CD38 on the surface of CAR T cells, the fratricide effect was reduced. This finding suggested that antibody blockade could abrogate CD38 target-driven fratricide. In summary, the addition of an antibody or a protein increased CAR T cell enrichment and expansion, and CD38 expression on the cell surface was preserved by blocking the CD38 Ag or CAR scFv domain. However, in the absence of the antibody or protein, antiCD38 CAR T cells did not expand well, and loss of CD38 Ag expression on the CAR T cell surface was observed (Fig. 3C). 2.4. Subsets of anti-CD38 CAR T cells It was previously reported that the lack of CAR T cells may be
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
369
Fig. 1. Construction of anti-CD38 CAR T cells and its expression. A: Schematic illustration for the lentiviral construct of anti-CD38 CAR T cells. The extracellular portion of the human CD38 receptor is a part of anti-CD38 CAR T cell, which was linked to transmembrane region and a CD8a hinge, followed by human 4-1BB (CD137) and CD3z intracellular signaling motif; not to scale. B and C: The expression of anti-CD38 CAR on T cells 15 days after transduction. MOCK T cells were constructed and used as control. Non-transduced T cells (NT) were used as negative control. D: Surface expression of CD38 in RPMI 8226 cell lines measured by flow cytometry in comparison with isotype control and K562 (CD38 negative control). E: The lysis ratio of anti-CD38 CAR Tcells on RPMI 8226 cells at different effector target (E:T) ratios for 24 h. F: The antitumor effects of anti-CD38 CAR T cells are shown in the histogram. G: The histogram showing that primary cells obtained from the patient's bone marrow were identified as CD38-positive cells by flow cytometry. H: Cell lysis ratio of CAR T cells to CD38 positive primary tumor cells at an E:T ratio of 1:1 for 6e48 h. Annexin V and 7AAD staining by flow cytometry was peformed. **P < 0.01.
due to “T cell fratricide” of anti-CD38-CAR T cells upon interaction with CD38 expressed on activated T cells (Alcantara et al., 2018). However, in the presence of the antibody or protein, CAR T cells were highly enriched during culture in vitro. To investigate the phenotype of the expanded CAR T cells, the cells were washed and analyzed by flow cytometry. CD62L and CD45RA are often used to distinguish among naïve T cells (CD62Lþ CD45RAþ; TN), terminally
differentiated T cells expressing CD45RA (CD62LeCD45RAþ; TD), effector memory T cells (CD62LeCD45RAe; TEM), and central memory T cells (CD62Lþ CD45RAe; TCM) (Golubovskaya and Wu, 2016; Caccamo et al., 2018). The MM27 group increased most in the relative frequency of naïve cells (defined as CD45RA and CD62L double-positive cells) (Fig. 4A), and the frequencies of CD279 (PD1) and CD366 (Tim-3) expressing cells were reduced most (Fig. 4B).
370
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 2. Expression of CD107a and CD38 antigen on anti-CD38 CAR T cells. A: Flow cytometry was performed to assess CD107a (LAMP-1), which was a potential marker in the process of degranulation. The CD107a on self-activated anti-CD38 CAR T cells has an expression ratio of 53.69% on day 6. MOCK-Ts was used as controls. B: CD38 antigen on anti-CD38 CAR T cells was almost completely lost at the same time. C: The cell debris of anti-CD38 CAR T cells as observed by phase contrast microscopy on day 6. Scale bar: 100 mm. D: Expansion of T cells in vitro for 15 days which was transduced with truncated anti-CD38 CAR lentiviruses. **P < 0.01.
These results suggested that the anti-CD38 CAR T cells were in a naïve stage and maintained a strong capacity for differentiation. In contrast, the group cultured without the antibodies or proteins expressed relatively higher levels of PD-1 and Tim-3 than the other groups. Interestingly, anti-CD38 CAR T cells survived and expanded as a predominant CD38eCD8þ T cell subset in the group treated with additives. Furthermore, antibody or protein blockade rescued the CD4þ subpopulation within the anti-CD38 CAR T cell population, suggesting that fratricide was dependent on a skewed CD4/CD8 ratio (Fig. 4C). The relative increase in the proportion of proliferating CD4þ T cells, as well as the elimination of fratricide effect between CD4þ T and CD8þ T cells during the manufacturing of antiCD38 CAR T cells, is the most likely explanation for the difference in the ratio.
2.5. Anti-CD38 CAR T cells effectively recognize and specifically lyse CD38-positive tumor cells For cytotoxicity assessment of anti-CD38 CAR T cells, we created the A549CD38þ cell line, which is derived from A549 cells and overexpressed CD38 (Fig. 5A). The expression of the mCherry protein in A549CD38þ cells was observed by fluorescence microscopy (Fig. 5B). CAR T cells were washed several times to eliminate the effect of residual antibodies or proteins in the medium. E:T ratios of 0.5:1, 1:1, and 5:1 were established with CAR T cells and target cells, and these cells were coincubated in cytokine-free medium for 6 h, 12 h, or 24 h. In addition, reactivity was measured by monitoring proinflammatory cytokine secretion. MOCK T cells were used at the same E:T ratios and A459 cells were employed as controls. As expected, compared to the control T cells,
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
371
significantly higher proportion of degranulated anti-CD38 CAR T cells than MOCK T cells upon exposure to target cells (Fig. 5G). Furthermore, the cytotoxicity mediated by anti-CD38 CAR T cells was dose and time dependent, and the CAR T cells exhibited increased cytotoxicity as the E:T ratio increased. The CD38-negative cell line A549 was not lysed by the anti-CD38 CAR T cells, and the growth of these cells was not inhibited by the MOCK T cells (Fig. 5C). Similar to the cytokine production results, the costimulated antibody or protein group demonstrated enhanced cytotoxicity compared to their unprotected counterparts. In contrast, no cytotoxicity was detected in the experiments with the A549 cell line, which lacks CD38 expression on the cell surface (Fig. 5D). Hence, we demonstrated that anti-CD38 CAR T cells possess potent cytotoxic activity against CD38-positive tumor cells. 2.6. Effective and persistent antitumor activity of anti-CD38 CAR T cells in xenograft tumors derived from RPMI 8226 cells Anti-CD38 CAR T cells have in vivo antitumor activity, which was evaluated by inoculating NOD/SCID/g-chain/ (NPG) mice with luciferase-labeled RPMI 8226 cells (Fig. 6A). When tumors were visible, the mice were treated with anti-CD38 CAR T cells using antibody (clone MM27). MOCK T cells and NaCl were used as controls. As shown in Fig. 6B, the anti-CD38 CAR T cells induced significant regression, or even elimination, of the RPMI 8226 tumors, while tumors continued to progress in the blank and MOCK T cell groups. Intriguingly, the anti-CD38 CAR T cells appeared to modestly delay tumor growth in the groups of mice receiving the anti-CD38 CAR T cells (Fig. 6C). On day 14 after T cell infusion, the anti-CD38 CAR T cells appeared in the peripheral blood (Fig. 6D). However, after 14e28 days of treatment, most of the mice in the NaCl and MOCK T cell-treated groups had died, indicating that the survival time of tumor-bearing mice was significantly prolonged after treatment with the anti-CD38 CAR T cells (Fig. 6E). In conclusion, costimulated anti-CD38 CAR T cells, a cellular immunotherapy modality, appeared to enhance the treatment of MM in vivo. 3. Discussion Fig. 3. Strategies to avoid CAR T cell lysis and to promote cell proliferation. A: Two distinct strategies were taken to avoid fratricide which was driven by targeting and to promote anti-CD38 CAR T cell production through blocking T cell surface CD38 Ag or CAR scfv by antibodies (Ab clone MM12T or clone MM27) or proteins (H02H or H08H). The total number of cells in each group during cell proliferation and enrichment was determined. B: The transduction efficiency in each group. C: The expression of CD38 on the cell surface in each group. *P < 0.05; **P < 0.01.
the anti-CD38 CAR T cells demonstrated significant cytotoxicity against A549CD38þ cells. As shown in Fig. 5C, the MM27 group exhibited higher average cytotoxic activity against A549 CD38þ cells (41.68% ± 3.02% at 6 h and 88.35% ± 7.02% at 24 h at an E:T ratio of 5:1) than the MOCK T cell group. When tumor cells were coincubated with CAR T cells or MOCK T cells at an E:T ratio of 1:1 for 24 h, the secreted cytokines were measured by ELISA. The levels of cytokines, including IL-2 (Fig. 5D), IFN-g (Fig. 5E) and TNF-a (Fig. 5F), were significantly higher in the supernatants of cocultures containing CAR T cells and A549CD38þ cells than in those of cocultures containing MOCK T cells. The MM27 group showed the slightly stronger cytokine secretion capacity than other antibody and protein groups tested. CD107a, which is expressed on the cell surface, has been described as a marker of cytotoxic CD8þ T cell degranulation, and its expression has been shown to be strongly upregulated on the cell surface in concordance with the loss of perforin following stimulation (Lorenzo-Herrero et al., 2019). In this study, we also found a
CAR T cell therapy represents a potent and targeted immunotherapeutic approach that has been used to treat patients with hematological malignancies. The toxicity profile of CAR T cell therapy is unique but not durable. Because CD19-negative recurrence often occurs after CART-19 treatment, there are two main reasons for treatment failure with CAR T cells: relapse with antigen loss and CAR T cell population loss (Wang et al., 2017). 3.1. CD38-based fratricide decreases the expansion and enrichment of anti-CD38 CAR T cells The human CD38 Ag is prevalently expressed by T lymphocytes, plasma cells, monocytes, NK cells and other cell types. The CD38 Ag is also overexpressed in various tumor cells, including those in BNHL, MM, and some relapsed and/or refractory diseases after CAR T cell therapy to eliminate CD38-positive cells (Chini et al., 2018). This antigen is considered an attractive target for treating hematological malignancies (Bannas and Koch-Nolte, 2018). Early studies confirmed that anti-CD38 CAR T cells have a strong antitumor ability. Mihara (Mihara et al., 2009) first reported that anti-CD38 CAR T cells self-lyse by fratricide. In contrast, Drent et al. (2016) speculated that no obvious dysfunction was observed in antiCD38 CAR T cells due to the lack of expression of CD38. Another study (An et al., 2018) observed some fratricide in new nanobodybased anti-CD38 CAR T cells. In this study, fratricide of the anti-
372
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 4. A detection of subsets of CAR T cells blocked with antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) at 15 days. A: Flow cytometry was performed to assess CD62L and CD45RA simultaneously, which were used to distinguish naïve (CD62Lþ CD45RAþ; TN), effector memory (CD62LeCD45RAe; TEM), central memory (CD62Lþ CD45RAe; TCM) and terminally differentiated T cell expressing CD45RA (CD62Le CD45RAþ; TD). B: Flow cytometry was performed to assess Tim-3 and PD-1 simultaneously, which were associated with T cell exhaustion/dysfunction. C: The expression levels of CD4 and CD8 were also measured by flow cytometry.
CD38 CAR T cells occurred, and the cells were unable to proliferate effectively because of the loss of CD38 expression. The most likely reason for this loss was that the potential target-driven fratricide of the anti-CD38 CAR T cells led to a “CAR T cell - autophagic and autostimulatory/enrichment” cycle.
3.2. Antibody- or protein-mediated blockade prevents anti-CD38 CAR T cell fratricide In the current study, we tried two distinct strategies to avoid fratricide driven by targeting CD38 and to increase anti-CD38 CAR T
Fig. 5. Efficacy and anti-tumor function of anti-CD38 CAR T cells in vitro. A: A549 was overexpressed with CD38 using a lentiviral vector with an mCherry tag and sorted by flow cytometry. B: The expression of mCherry protein was detected by fluorescence microscopy to show the expression of CD38 in A549CD38þ cells (original magnification, 100). C: A549 and A549CD38þ cells were cocultured with anti-CD38 CAR T cells at effector-to-target ratios of 0.5:1, 1:1, or 5:1 for 6 h, 12 h, and 24 h. The lysis of A549CD38þ cells by anti-CD38 CAR T cells was observed at each time point. Negative effector was A459 cells, as well as target cell controls. D-F: When co-incubating the tumor cells with anti-CD38 CAR T cells or MOCK T cells at an E: T ratio of 1:1 for 24 h, the secreted cytokines were determined by ELISA, including IL-2 (D), IFN-g (E), and TNF-a (F). G: Flow cytometry was carried to detect the expression levels of CD107a in anti-CD38 CAR T cells or MOCK T cells. *P < 0.05, **P < 0.01.
374
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 6. Strong antitumor activity of anti-CD38 CAR T cells in an MM model. A: Schematic of this study. NPG mice were inoculated intraperitoneally (i.p.) in the flank with 1 106 RPMI 8226-mCherry-luc cells on day 7. After 1 week, anti-CD38 CAR T cells treated by antibody (clone MM27) were injected via the tail vein on day 0. MOCK-Ts and NaCl were used as controls. B: Bioluminescence imaging (BLI) in each mouse was measured to study the tumor burden, and the changes of tumor burden within 6 weeks after CAR-T reinfusion were presented. C: ROI of each BLI at day 14 was analyzed. Error bars denote the s.e.m., and two-way ANOVA was chosen to compare the results. **P < 0.01. D: T cell percentage at day 14 in the peripheral blood of mice intraperitoneally injected with RPMI 8226. Unpaired t-test was used for comparison of results. **P < 0.01. E: Survival curve of mice intraperitoneally injected with RPMI 8226 cells. Results were analyzing using a Log-rank (Mantel-Cox) test. **P < 0.01.
cell production by blocking the T cell surface expression of CD38 or the CAR scFv domain with an antibody (clone MM12T or clone MM27) or a protein (H02H or H08H). The results showed that the antibodies and proteins impacted CAR T cell fratricide and
promoted CAR T cell expansion and enrichment. Furthermore, the addition of the antibody clone MM27 was most effective in improving CAR T cell yield, increasing the relative frequency of naïve cells (defined as CD62L and CD45RA double-positive cells)
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
and reducing the frequencies of CD279 (PD-1) (Catakovic et al., 2017) and CD366 (Tim-3) expressing cells (Banerjee and Kane, 2018). This result suggested that the treated anti-CD38 CAR T cells had improved capacities for self-renewal and long-term survival. These data have powerful implications on the large-scale expansion of CAR T cells after tumor cell recognition. In addition, both protein treatments generated comparable anti-CD38 CAR T cell populations. Furthermore, these methods have been used in clinical trials to efficiently and reproducibly produce anti-CD38 CAR T cells. Previous preclinical studies have indicated that differences in CAR T efficacy depend on the different populations of CD4þ T cells and CD8þ T cells. The manufacture of CAR T cell products with a defined 1:1 ratio of CD4þ/CD8þ T cells was demonstrated to be the most efficacious approach (Sommermeyer et al., 2016). In this study, CD4þ anti-CD38 CAR T cells suffered severe lysis in the absence of antibodies or proteins protection during culture. In addition, a subset of CD8þ CAR T cells expanded and proliferated. The lack of helper CD4þ T cells might reduce the antitumor ability and persistence of generated CAR T cells. Conversely, CD4þ CAR T cells were rescued by pretreatment with antibodies or proteins during CAR T cell culture. These results indicate that fratricide is dependent on the CD4þ/CD8þ T cell ratio. Differences in this ratio are most likely due to the proliferation of CD4þ T cells or the reduction in CD4þ T cell numbers caused by CD8þ T cells. 3.3. Specific cytotoxicity of anti-CD38 CAR T cells against CD38þ cell lines In the current study, we constructed anti-CD38 CAR T cells and evaluated their antitumor activity and persistence. The anti-CD38 CAR T cells were cocultured with CD38þ tumor cell lines to assay cytolysis. It has consistently been demonstrated by in vitro results that CD38þ cells can be lysed efficiently and specifically by antiCD38 CAR T cells in a dose- and time-dependent manner. Similarly, the antitumor effect of CAR T cells on primary CD38þ tumor cells has also been demonstrated. The antitumor effect of CAR T cells is positively correlated with the secretion levels of various cytokines, and the levels of the proinflammatory factors IL-2, IFN-g and TNF-a were significantly upregulated in the supernatants of A549CD38þ cells. Cell surface-expressed CD107a is considered to be a marker of CD8þ T cell degranulation. In this experiment, we found a significantly higher proportion of degranulated anti-CD38 CAR T cells than degranulated MOCK T cells upon exposure to target cells. Similar antitumor activity was observed in a NOD/SCID/g-chain/ (NPG) mouse xenograft model. Specificity was confirmed by using MOCK T cells and A549 cells as negative controls. In conclusion, targeting CD38 with CAR T cells is an enticing and novel approach to further improve antitumor effects. However, the target-driven fratricide of anti-CD38 CAR T cells remains a major problem. Our strategy avoided cell lysis and obtained a sufficient preparation of cells for CAR T cell manufacturing, especially in the presence of the antibody clone MM27. We will verify this strategy in future experiments. Anti-CD38 CAR T cells still face significant challenges in future clinical studies. Since the CD38 Ag is widely expressed in normal cells and tissues, there is a risk of infused CAR T cells attacking these cells and tissues. Most importantly, in myeloid progenitors, CD38 is a common differentiation antigen (Miyawaki et al., 2017), and abnormal hematopoiesis may result after targeting the CD38 Ag. In this study, immunotherapy mediated by anti-CD38 CAR T cells has the potential to eliminate CD38þ tumor cells and overcome target-driven fratricide of CAR T cells through blockade of T cell surface-expressed CD38 Ag or CAR scFv by antibodies or proteins. Anti-CD38 CAR T cells demonstrated specific and strong antitumor
375
effects in vitro and in vivo. The results will help to further enhance the expansion, persistence and function of anti-CD38 CAR T cells in subsequent clinical trials (NCT03754764). 4. Methods and materials 4.1. Cell lines and culture reagents The human MM cell line RPMI 8226 (CD38-positive) was used in immune-based assays. MM cells were cultured in RPMI medium (Sigma-Aldrich, St. Louis, MO, USA) containing 10% fetal calf serum (Atlas Biologicals, USA). The human lung adenocarcinoma cell line A549 was used as the antigen-negative control. CD38overexpressing A549 cells were constructed with a lentiviral vector with an mCherry tag and named A549CD38þ cells. The immortalized normal fetal renal 293T cell line was cultured in DMEM for lentiviral packaging. CD38-positive primary tumor cells were obtained from a patient. Complete medium was used for the culture of all cell lines: 100 mg/mL streptomycin sulfate, 100 U/mL penicillin and 10% heat inactivated fetal bovine serum (FBS) were added to supplemented RPMI-1640 medium. CD38 proteins (H02H and H08H) and antibodies (clone MM12T and MM27) were purchased from Beijing Sino Biological Co., Ltd. 4.2. Constructing anti-CD38 CAR T cells and producing lentiviruses The anti-CD38 CAR consisted of the human anti-CD38 scFv followed by the intracellular signaling motifs of human CD137 and CD3z (Fig. 2A), which have been described previously (Wang et al., 2015; Zhang et al., 2016). The pWPT-anti-CD38 CAR plasmid and ps-PAX2 packaging plasmid were transfected into 293 T cells. All the lentiviruses in this study were obtained from concentrated stocks. Before experimentation or titration, viruses were aliquoted and stored at 80 C. 4.3. Transduction of human T cells Primary human T cells were isolated from healthy normal donors. Peripheral blood mononuclear cells (PBMCs) were suspended in XeVIVO 15 medium directly (Lonza, USA) with an anti-CD3 monoclonal antibody (mAb; 500 ng/mL) and recombinant human IL-2 (300 U/mL) (PeproTech, USA). Transduction mediated by a lentivirus was performed on day 0. Briefly, in the presence of protamine sulfate (Sigma-Aldrich) at a concentration of 10 mg/mL, 1 106 T cells were infected with lentiviral vectors encoding CAR constructs overnight. After transduction, the cultures were transferred to fresh XeVIVO 15 medium supplemented with IL-2 addition every other day, and the cell density was maintained at 0.5e1 106 cells/mL. 4.4. Antibodies or proteins inhibition assays Two distinct strategies were used to avoid target-driven fratricide. The first strategy was to use the mouse IgG1-type antibody clones MM12T or MM27 to block the anti-CD38 CAR on the T cell surface. The second strategy was to use the protein H02H or H08H to block the cell-transduced scFv domain of the CAR. In the process of cell culture, the appropriate protein or antibody was added to the CAR T cell culture medium at the end of cell transduction. MOCK T cell and no additive CAR T cell groups were established at the same time. In coculture experiments, to eliminate the effects of residual antibody or protein, CAR T cells were washed several times in medium.
376
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
4.5. Flow cytometric analysis The following fluorochrome-conjugated antibodies purchased from BD (Biosciences, USA) were used in this study: APC-Cy7 antihuman CD3, FITC anti-human CD4, PE anti-human CD8, PE antihuman CD366 (Tim-3). APC anti-human CD269 (PD-1), APC antihuman CD45RA, PE anti-human CD62L, and APC anti-human CD107a. Annexin V-PE and 7-Aminoactinomycin D (7-AAD) were used for viability assessment (Crowley et al., 2016). In experiments with the addition of the mouse IgG1-type antibody clones MM12T and MM27, a goat anti-mouse IgG1 antibody (FITC) was chosen to analyze CD38 expression on cells. In the remaining groups, including the MOCK T cells with the addition of protein H02H or H08H groups and the no additive CAR T cell groups, the CAR T cells were stained with a PE-conjugated anti-human CD38 antibody. A recombinant CD38-Fc protein and a goat anti-mouse Fc (APC) antibody were used to evaluate surface expression. All analyses used matched secondary and isotype antibodies. Biotin-spaffinipure goat anti-mouse IgG F(ab')2 fragment and a PEsreptavidin antibody were used to detect MOCK T cells in the following experiments. A Beckman Coulter CytoFlex flow cytometer was used to perform flow cytometry, and FlowJo software was used to analyze the data (Version 10. 0. 7, FlowJo, Ashland, OR, USA). 4.6. Cytokine release assays Anti-CD38 CAR T cells and target cells were cocultured in a 1:1 ratio, followed by evaluation with a cytokine release assay. The cells were cultured in 96-well plates, with 200 mL of RPMI-1640 medium in each well. When the experiment ended, an ELISA (BioLegend, USA) was performed using cell-free coculture supernatants to measure the secretion of TNF-a, IL-2 and IFN-g. The results were measured three times and were expressed as an average. 4.7. Cytotoxicity assays Adherent A549 and A549CD38þ cells in each of the above mentioned groups were cultured in a flat-bottomed 96-well plate for 20 h at E:T ratios of 0.5:1, 1:1, and 5:1 for 6 h, 12 h, or 24 h. AntiCD38 CAR T cells were washed, and the remaining adherent A549CD38þ cells were labeled with alamar Blue (ThermoFisher Scientific, USA) in serum-free medium (Xu et al., 2015; Breman et al., 2018) for 4 h. The Infinite M200 Pro (TECAN, Switzerland) was used to measure viable cells at 530 nm, and then the relative cytolytic activity was calculated. 4.8. Xenograft model of MM To establish a xenotransplanted tumor model, female NPG mice (6e8 weeks old) were obtained from Beijing Vitalstar Biotechnology Co., Ltd. and inoculated intraperitoneally (i.p.) in the flank with 1 106 RPMI 8226-mCherry-luc cells on day 7. The tumors became palpable at approximately 1 week, and the mice were injected via the tail vein with anti-CD38 CAR T cells or MOCK T cells (1 106 cells suspended in 100 mL of PBS) on day 0. Bioluminescence imaging (BLI) was used to monitor tumor growth. We performed the animal experiments according to the Animal Protection Guidelines of the People's Liberation Army General Hospital (PLAGH). 4.9. Statistical analysis All experiments were performed three times, and the results are presented as the mean ± SD. Differences among groups were examined using one-way analysis of variance (ANOVA). Differences
in the number of transferred T cells were evaluated by Student's ttest. Analyses were performed with IBM SPSS 23 software (IBM, USA), and a two-tailed P < 0. 01 was considered statistically significant. Acknowledgments We thank all the patients for consenting to provide tissue samples. This work was supported by the grants from the National Natural Science Foundation of China (Nos. 81830002 and 31870873 to W.D.H.) and the National Key Research and Development Program of China (Nos. 2016YFC1303501 and 2016YFC1303504 to WDH, and No. 2017YFC0909803 to W.Y.). References Alcantara, M., Tesio, M., June, C.H., Houot, R., 2018. CAR T-cells for T-cell malignancies: challenges in distinguishing between therapeutic, normal, and neoplastic T-cells. Leukemia 32, 2307e2315. An, N., Hou, Y.N., Zhang, Q.X., Li, T., Zhang, Q.L., Fang, C., Chen, H., Lee, H.C., Zhao, Y.J., Du, X., 2018. Anti-multiple myeloma activity of nanobody-based anti-CD38 chimeric antigen receptor T cells. Mol. Pharm. 15, 4577e4588. Atanackovic, D., Steinbach, M., Radhakrishnan, S.V., Luetkens, T., 2016. Immunotherapies targeting CD38 in multiple myeloma. OncoImmunology 5 e1217374. Banerjee, H., Kane, L.P., 2018. Immune Regulation by Tim-3. F1000Res, vol. 7, p. 316. Bannas, P., Koch-Nolte, F., 2018. Perspectives for the development of CD38-Specific heavy chain antibodies as therapeutics for multiple myeloma. Front. Immunol. 9, 2559. Blatt, K., Menzl, I., Eisenwort, G., Cerny-Reiterer, S., Herrmann, H., Herndlhofer, S., Stefanzl, G., Sadovnik, I., Berger, D., Keller, A., Hauswirth, A., Hoermann, G., Willmann, M., Rulicke, T., Sill, H., Sperr, W.R., Mannhalter, C., Melo, J.V., Jager, U., Sexl, V., Valent, P., 2018. Phenotyping and target expression profiling of CD34þ/ CD38- and CD34þ/CD38þ stem- and progenitor cells in acute lymphoblastic leukemia. Neoplasia 20, 632e642. Bonello, F., D'Agostino, M., Moscvin, M., Cerrato, C., Boccadoro, M., Gay, F., 2018. CD38 as an immunotherapeutic target in multiple myeloma. Expert Opin. Biol. Ther. 18, 1209e1221. Breman, E., Demoulin, B., Agaugue, S., Mauen, S., Michaux, A., Springuel, L., Houssa, J., Huberty, F., Jacques-Hespel, C., Marchand, C., Marijsse, J., Nguyen, T., Ramelot, N., Violle, B., Daro, D., De Waele, P., Gilham, D.E., Steenwinckel, V., 2018. Overcoming target driven fratricide for T cell therapy. Front. Immunol. 9, 2940. Brudno, J.N., Kochenderfer, J.N., 2019. Recent advances in CAR T-cell toxicity: mechanisms, manifestations and management. Blood Rev. 34, 45e55. Caccamo, N., Joosten, S.A., Ottenhoff, T.H.M., Dieli, F., 2018. Atypical human effector/ memory CD4þ T cells with a naive-like phenotype. Front. Immunol. 9, 2832. Catakovic, K., Klieser, E., Neureiter, D., Geisberger, R., 2017. T cell exhaustion: from pathophysiological basics to tumor immunotherapy. Cell Commun. Signal. 15, 1. Chehab, S., Panjic, E.H., Gleason, C., Lonial, S., Nooka, A.K., 2018. Daratumumab and its use in the treatment of relapsed and/or refractory multiple myeloma. Future Oncol. 14, 3111e3121. Chini, E.N., Chini, C.C.S., Espindola Netto, J.M., de Oliveira, G.C., van Schooten, W., 2018. The pharmacology of CD38/NADase: an emerging target in cancer and diseases of aging. Trends Pharmacol. Sci. 39, 424e436. Crowley, L.C., Marfell, B.J., Scott, A.P., Waterhouse, N.J., 2016. Quantitation of apoptosis and necrosis by annexin V binding, propidium iodide uptake, and flow cytometry. Cold Spring Harb. Protoc. 2016 https://doi.org/10.1101/pdb. prot087288. Drent, E., Groen, R.W., Noort, W.A., Themeli, M., Lammerts van Bueren, J.J., Parren, P.W., Kuball, J., Sebestyen, Z., Yuan, H., de Bruijn, J., van de Donk, N.W., Martens, A.C., Lokhorst, H.M., Mutis, T., 2016. Pre-clinical evaluation of CD38 chimeric antigen receptor engineered T cells for the treatment of multiple myeloma. Haematologica 101, 616e625. Feng, K.C., Guo, Y.L., Liu, Y., Dai, H.R., Wang, Y., Lv, H.Y., Huang, J.H., Yang, Q.M., Han, W.D., 2017. Cocktail treatment with EGFR-specific and CD133-specific chimeric antigen receptor-modified T cells in a patient with advanced cholangiocarcinoma. J. Hematol. Oncol. 10, 4. Golubovskaya, V., Wu, L., 2016. Different subsets of T cells, memory, effector functions, and CAR-T immunotherapy. Cancers (Basel) 8, 36. Han, X., Wang, Y., Han, W.D., 2018. Chimeric antigen receptor modified T-cells for cancer treatment. Chronic Dis. Transl. Med. 4, 225e243. Kloss, S., Oberschmidt, O., Dahlke, J., Vu, X.K., Neudoerfl, C., Kloos, A., Gardlowski, T., Matthies, N., Heuser, M., Meyer, J., Sauer, M., Falk, C., Koehl, U., Schambach, A., Morgan, M.A., 2019. Preclinical assessment of suitable natural killer (NK) cell sources for chimeric antigen receptor (CAR)-NK-based "off-the-shelf" AML immunotherapies. Hum. Gene Ther. 30. https://doi.org/10.1089/hum.2018.247. Lorenzo-Herrero, S., Sordo-Bahamonde, C., Gonzalez, S., Lopez-Soto, A., 2019. CD107a degranulation assay to evaluate immune cell antitumor activity. Methods Mol. Biol. 1884, 119e130. McHayleh, W., Bedi, P., Sehgal, R., Solh, M., 2019. Chimeric antigen receptor T-cells:
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377 the future is now. J. Clin. Med. 8 pii: E207. Mele, S., Devereux, S., Pepper, A.G., Infante, E., Ridley, A.J., 2018. Calcium-RasGRP2Rap1 signaling mediates CD38-induced migration of chronic lymphocytic leukemia cells. Blood Adv. 2, 1551e1561. Mihara, K., Bhattacharyya, J., Kitanaka, A., Yanagihara, K., Kubo, T., Takei, Y., Asaoku, H., Takihara, Y., Kimura, A., 2012. T-cell immunotherapy with a chimeric receptor against CD38 is effective in eliminating myeloma cells. Leukemia 26, 365e367. Mihara, K., Yanagihara, K., Takigahira, M., Imai, C., Kitanaka, A., Takihara, Y., Kimura, A., 2009. Activated T-cell-mediated immunotherapy with a chimeric receptor against CD38 in B-cell non-Hodgkin lymphoma. J. Immunother. 32, 737e743. Mihara, K., Yanagihara, K., Takigahira, M., Kitanaka, A., Imai, C., Bhattacharyya, J., Kubo, T., Takei, Y., Yasunaga, S., Takihara, Y., Kimura, A., 2010. Synergistic and persistent effect of T-cell immunotherapy with anti-CD19 or anti-CD38 chimeric receptor in conjunction with rituximab on B-cell non-Hodgkin lymphoma. Br. J. Haematol. 151, 37e46. Miyawaki, K., Iwasaki, H., Jiromaru, T., Kusumoto, H., Yurino, A., Sugio, T., Uehara, Y., Odawara, J., Daitoku, S., Kunisaki, Y., Mori, Y., Arinobu, Y., Tsuzuki, H., Kikushige, Y., Iino, T., Kato, K., Takenaka, K., Miyamoto, T., Maeda, T., Akashi, K., 2017. Identification of unipotent megakaryocyte progenitors in human hematopoiesis. Blood 129, 3332e3343. Morandi, F., Horenstein, A.L., Costa, F., Giuliani, N., Pistoia, V., Malavasi, F., 2018. CD38: a target for immunotherapeutic approaches in multiple myeloma. Front. Immunol. 9, 2722. Naik, J., Themeli, M., de Jong-Korlaar, R., Ruiter, R.W.J., Poddighe, P.J., Yuan, H., de Bruijn, J.D., Ossenkoppele, G.J., Zweegman, S., Smit, L., Mutis, T., Martens, A.C.M., van de Donk, N., Groen, R.W.J., 2018. CD38 as a therapeutic target for adult acute myeloid leukemia and T-cell acute lymphoblastic leukemia. Haematologica 104, e100ee103. Pick, M., Vainstein, V., Goldschmidt, N., Lavie, D., Libster, D., Gural, A., Grisariu, S., Avni, B., Ben Yehuda, D., Gatt, M.E., 2018. Daratumumab resistance is frequent in advanced-stage multiple myeloma patients irrespective of CD38 expression and is related to dismal prognosis. Eur. J. Haematol. 100, 494e501.
377
Plesner, T., Krejcik, J., 2018. Daratumumab for the treatment of multiple myeloma. Front. Immunol. 9, 1228. Roddie, C., O'Reilly, M., Dias Alves Pinto, J., Vispute, K., Lowdell, M., 2019. Manufacturing chimeric antigen receptor T cells: issues and challenges. Cytotherapy 21, 327e340. Sommermeyer, D., Hudecek, M., Kosasih, P.L., Gogishvili, T., Maloney, D.G., Turtle, C.J., Riddell, S.R., 2016. Chimeric antigen receptor-modified T cells derived from defined CD8þ and CD4þ subsets confer superior antitumor reactivity in vivo. Leukemia 30, 492e500. van de Donk, N., 2018. Immunomodulatory effects of CD38-targeting antibodies. Immunol. Lett. 199, 16e22. van de Donk, N., Usmani, S.Z., 2018. CD38 antibodies in multiple myeloma: mechanisms of action and modes of resistance. Front. Immunol. 9, 2134. Wang, J., Hu, Y., Huang, H., 2017. Acute lymphoblastic leukemia relapse after CD19targeted chimeric antigen receptor T cell therapy. J. Leukoc. Biol. 102, 1347e1356. Wang, Q.S., Wang, Y., Lv, H.Y., Han, Q.W., Fan, H., Guo, B., Wang, L.L., Han, W.D., 2015. Treatment of CD33-directed chimeric antigen receptor-modified T cells in one patient with relapsed and refractory acute myeloid leukemia. Mol. Ther. 23, 184e191. Xu, M., McCanna, D.J., Sivak, J.G., 2015. Use of the viability reagent prestoplue in comparison with alamarblue and MTT to assess the viability of human corneal epithelial cells. J. Pharmacol. Toxicol. Methods 71, 1e7. Zaja, F., Tabanelli, V., Agostinelli, C., Calleri, A., Chiappella, A., Varettoni, M., Luigi Zinzani, P., Volpetti, S., Sabattini, E., Fanin, R., Pileri, S.A., 2017. CD38, BCL-2, PD1, and PD-1L expression in nodal peripheral T-cell lymphoma: possible biomarkers for novel targeted therapies? Am. J. Hematol. 92, E1eE2. Zhang, C., Kasi, A., 2018. Chimeric Antigen Receptor (CAR) T-Cell Therapy. StatPearls, Treasure Island (FL). Zhang, W.Y., Wang, Y., Guo, Y.L., Dai, H.R., Yang, Q.M., Zhang, Y.J., Zhang, Y., Chen, M.X., Wang, C.M., Feng, K.C., Li, S.X., Liu, Y., Shi, F.X., Luo, C., Han, W.D., 2016. Treatment of CD20-directed chimeric antigen receptor-modified T cells in patients with relapsed or refractory B-cell non-Hodgkin lymphoma: an early phase IIa trial report. Signal Transduct. Target. Ther. 1, 16002.
Contents lists available at ScienceDirect
Journal of Genetics and Genomics Journal homepage: www.journals.elsevier.com/journal-of-geneticsand-genomics/
Original research
Blocking CD38-driven fratricide among T cells enables effective antitumor activity by CD38-specific chimeric antigen receptor T cells Zhitao Gao, Chuan Tong, Yao Wang, Deyun Chen, Zhiqiang Wu*, Weidong Han* Department of Molecular Biology, Institute of Basic Medicine, School of Life Sciences, Medical School of Chinese PLA, Chinese PLA General Hospital, Beijing, 100086, China
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 April 2019 Received in revised form 9 June 2019 Accepted 17 June 2019 Available online 13 August 2019
Chimeric antigen receptor T-cell (CAR T) therapy is a kind of effective cancer immunotherapy. However, designing CARs remains a challenge because many targetable antigens are shared by T cells and tumor cells. This shared expression of antigens can cause CAR T cell fratricide. CD38-targeting approaches (e.g., daratumumab) have been used in clinical therapy and have shown promising results. CD38 is a kind of surface glycoprotein present in a variety of cells, such as T lymphocytes and tumor cells. It was previously reported that CD38-based CAR T cells may undergo apoptosis or T cell-mediated killing (fratricide) during cell manufacturing. In this study, a CAR containing a sequence targeting human CD38 was designed to be functional. To avoid fratricide driven by CD38 and ensure the production of CAR T cells, two distinct strategies based on antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) were used to block CD38 or the CAR single-chain variable fragment (scFv) domain, respectively, on the T cell surface. The results indicated that the antibodies or proteins, especially the antibody MM27, could affect CAR T cells by inhibiting fratricide while promoting expansion and enrichment. Anti-CD38 CAR T cells exhibited robust and specific cytotoxicity to CD38þ cell lines and tumor cells. Furthermore, the levels of the proinflammatory factors TNF-a, IFN-g and IL-2 were significantly upregulated in the supernatants of A549CD38þ cells. Finally, significant control of disease progression was demonstrated in xenograft mouse models. In conclusion, these findings will help to further enhance the expansion, persistence and function of anti-CD38 CAR T cells in subsequent clinical trials. Copyright © 2019, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, and Genetics Society of China. Published by Elsevier Limited and Science Press. All rights reserved.
Keywords: Chimeric antigen receptor CD38 T cells Immunotherapy CD38 antibody
1. Introduction Adoptive cell therapy (ACT) with genetically modified T cells expressing a chimeric antigen receptor (CAR), which are artificially engineered receptors expressed on the cell surface of T cells with non-HLA-restricted tumor antigens, allows CARs to activate T cells and specifically guide them to tumor cells to perform their function (Han et al., 2018; Zhang and Kasi, 2018). However, designing CARs remains a challenge because of the shared expression of many targetable antigens between normal cells and tumor cells, including targets expressed on the T cell surface, such as NKG2D, CS1, CD7, and CD5. This shared expression of antigens can cause fratricide of CAR T cells, inhibiting their proliferation and viability
* Corresponding authors. E-mail addresses: [email protected] (Z. Wu), [email protected] (W. Han).
and causing unpredictable on-target, off-tumor toxicity in clinical studies (Brudno and Kochenderfer, 2019). Human CD38 antigen (45 kDa) is a type II transmembrane glycoprotein that is expressed by monocytes, plasma cells and immature cells and activates T and B lymphocytes (Atanackovic et al., 2016; Bonello et al., 2018; Morandi et al., 2018). In the normal hematopoietic system, CD38 is expressed only by precursors that are “lineage committed” (Mihara et al., 2012) and is rarely expressed by multipotent stem cells and nonhematopoietic tissues. However, it is widely expressed on various types of cancer cells. CD38 is overexpressed in several lymphocyte malignancies, including B-cell non-Hodgkin lymphoma (B-NHL) (Mihara et al., 2010), multiple myeloma (MM), and collagen disease. CD38 is also expressed in some refractory and/or relapsed acute lymphoblastic leukemias (ALL) (Blatt et al., 2018; Naik et al., 2018), chronic lymphocytic leukemia (CLL) (Mele et al., 2018) and non-Hodgkin lymphoma (NHL) associated with CD38-positive cells (Zaja et al., 2017), which are especially associated with relapse and/or
https://doi.org/10.1016/j.jgg.2019.06.007 1673-8527/Copyright © 2019, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, and Genetics Society of China. Published by Elsevier Limited and Science Press. All rights reserved.
368
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
refractory cancer after anti-CD19 CAR T cell therapy. Furthermore, sustained expression of CD38 on tumor cells is associated with a poor prognosis (Pick et al., 2018). On the basis of previous studies, CD38 is regarded as an attractive target, and targeting CD38 is a potential treatment strategy for hematological malignancies. An anti-CD38 antibody (daratumumab) has been approved for the treatment of MM by the Food and Drug Administration (FDA) (Chehab et al., 2018; van de Donk and Usmani, 2018), and preclinical and clinical studies have demonstrated the efficacy and safety of the anti-CD38 antibody (Plesner and Krejcik, 2018; van de Donk, 2018). CD38 plays an important role in CAR T cell immunotherapy as a molecular target. However, CD38 is also present on the surface of NK cells and T cells. Thus, these cells may undergo suicide upon the transduction of CAR T cells. In this study, a lentiviral vector carrying a CAR comprising an anti-CD38 single-chain variable fragment (scFv) domains was designed and the antitumor activity of antiCD38 CAR-modified T cells was evaluated. As previously reported, activated T cells can also express CD38, suggesting that these T cells may undergo apoptosis or T cell-mediated killing (fratricide) during cell manufacturing (McHayleh et al., 2019; Roddie et al., 2019). To inhibit target-driven fratricide, two different strategies based on CD38 proteins or blocking antibodies were investigated. It was found that antibodies or proteins can affect anti-CD38 CAR T cells by inhibiting fratricide while promoting expansion and enrichment. Further, human T cells have been demonstrated to be resistant to CD38-positive cells with high efficiency both in vivo and in vitro in the present of anti-CD38 CAR T cells. These strategies have been incorporated into on-going clinical trials (NCT03754764) testing the efficacy of this CAR-modified T-cell immunotherapy for the treatment of relapsed and/or refractory leukemia associated with CD38-positive cells, especially for the treatment of relapsed and/or refractory disease after CAR T cell therapy.
2. Results 2.1. An anti-CD38 CAR design and the expression of the CAR on the surface of T cells To test whether normal T cells can recognize tumor cells via a CD38-specific CAR, we created a CAR construct with the human anti-CD38 scFv linked to a CD8a hinge and transmembrane region followed by the human 4-1BB (CD137) and CD3z intracellular signaling motifs, as shown in Fig. 1A. Similarly, MOCK T cells were constructed and used as a control. HEK-293T cells cotransfected with packaging plasmids were used to prepare lentiviral particles encoding the CAR plasmid (Feng et al., 2017). To confirm anti-CD38 CAR expression in transduced T cells, a CD38 Fc-tagged protein was used, followed by incubation with a goat anti-Fc antibody conjugated with APC that cross-reacts with the anti-CD38 CAR. The expression of anti-CD38 CAR was then analyzed by flow cytometry. The anti-CD38 CAR T cell transduction efficiency reached 90.53% ± 8.72% after 15 days (Fig. 1B and C). To assess cytotoxicity, we cocultured anti-CD38 CAR T cells with CD38-positive RPMI 8226 cells (Fig. 1D). After 24 h, 62.40% ± 6.73% of the RPMI 8226 cells were lysed by the anti-CD38 CAR T cells (Fig. 1E and F) at an effector-to-target (E:T) ratio of 5:1, while little lysis was observed in the coculture with MOCK T cells. In contrast, no cytotoxicity was detected in a K562 cell assay, as K562 cells lack CD38 expression. In an assay with CD38-positive primary tumor cells (Fig. 1G), anti-CD38 CAR T cells were incubated with the primary cells at an E:T ratio of 1:1 for 6e48 h. The efficiency of tumor cell lysis by the CAR T cells reached 22.76% at 24 h and 40.44% at 48 h (Fig. 1H).
2.2. CD38-driven fratricide of anti-CD38 CAR T cells To confirm the occurrence of cytotoxicity via CAR T cell fratricide, flow cytometry was carried out to detect the expression of CD107a during anti-CD38 CAR T cell expansion over 6 days (Fig. 2A). Normally, CD107a is regarded as a marker of CAR T cell degranulation. Once CAR T cells contact their target antigen, they are rapidly activated, express CD107a, and undergo degranulation (Kloss et al., 2019). However, human CD38 antigen (Ag) was prevalently expressed on T lymphocytes and caused CAR T cell activation. Cytotoxicity increased with increasing expression of CD107a, which may be associated with the loss of the CD38 Ag from the cell surface (Fig. 2B), and many dead cells and much debris were simultaneously observed (Fig. 2C). The decrease in CD38 expression on CAR T cells or the lysis of CD38þ CAR T cells by activated anti-CD38 CAR T cells appears to lead to a “CAR T cell - autophagic and autostimulatory/enrichment” cycle. In initial experiments, the population of anti-CD38 CAR T cells was measured during the cell expansion process. We found that compared with MOCK T cells, anti-CD38 CAR T cells could not achieve a rapid expansion ratio. Although T cells expressing this CAR have been successfully generated, there were significant differences in enrichment and proliferation between the CAR T cell group and the MOCK T cell group (Fig. 2D). We hypothesized that target-driven fratricide caused the loss of cell viability among the anti-CD38 CAR T cells. 2.3. Antibodies or proteins promote the enrichment and expansion of CAR T cells To avoid fratricide driven by targeting and to improve anti-CD38 CAR T cell production for clinical application, two distinct methods were investigated. The first approach focused on CD38 proteins to block the anti-CD38 CAR scFv. The second strategy involved antibody blockade of the CD38 Ag during CAR T cell production. Therefore, we tested whether an antibody or a protein could abrogate anti-CD38 CAR T cell-mediated fratricide during T cell culture. Antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) were added during the culture of anti-CD38 CAR T cells, and MOCK T cell and no additive CAR T cell groups were established at the same time. After 15 days of cell culture and expansion, the anti-CD38 CAR T cell transduction efficiency ranged from 58.61% to 94%, and the mean MOCK T cell transduction efficiency reached 83.57% ± 6.57%. In particular, the total cell number of the MM27 group was highly enriched ((102.36 ± 10.98) 106), and the transduction efficiency reached 76.49% ± 4.83% (Fig. 3A and B). Similarly, the total number of MM12T group reached (87.64 ± 5.79) 106, and the transduction efficiency reached 67.77% ± 5.16%. Furthermore, compared with the no additive CAR T cell group, the two groups treated with a protein (H02H or H08H) to block the CAR scFv domain also exhibited significantly improved CAR T cell expansion and transduction efficiency. Since the antibodies used blocked CD38 on the surface of CAR T cells, the fratricide effect was reduced. This finding suggested that antibody blockade could abrogate CD38 target-driven fratricide. In summary, the addition of an antibody or a protein increased CAR T cell enrichment and expansion, and CD38 expression on the cell surface was preserved by blocking the CD38 Ag or CAR scFv domain. However, in the absence of the antibody or protein, antiCD38 CAR T cells did not expand well, and loss of CD38 Ag expression on the CAR T cell surface was observed (Fig. 3C). 2.4. Subsets of anti-CD38 CAR T cells It was previously reported that the lack of CAR T cells may be
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
369
Fig. 1. Construction of anti-CD38 CAR T cells and its expression. A: Schematic illustration for the lentiviral construct of anti-CD38 CAR T cells. The extracellular portion of the human CD38 receptor is a part of anti-CD38 CAR T cell, which was linked to transmembrane region and a CD8a hinge, followed by human 4-1BB (CD137) and CD3z intracellular signaling motif; not to scale. B and C: The expression of anti-CD38 CAR on T cells 15 days after transduction. MOCK T cells were constructed and used as control. Non-transduced T cells (NT) were used as negative control. D: Surface expression of CD38 in RPMI 8226 cell lines measured by flow cytometry in comparison with isotype control and K562 (CD38 negative control). E: The lysis ratio of anti-CD38 CAR Tcells on RPMI 8226 cells at different effector target (E:T) ratios for 24 h. F: The antitumor effects of anti-CD38 CAR T cells are shown in the histogram. G: The histogram showing that primary cells obtained from the patient's bone marrow were identified as CD38-positive cells by flow cytometry. H: Cell lysis ratio of CAR T cells to CD38 positive primary tumor cells at an E:T ratio of 1:1 for 6e48 h. Annexin V and 7AAD staining by flow cytometry was peformed. **P < 0.01.
due to “T cell fratricide” of anti-CD38-CAR T cells upon interaction with CD38 expressed on activated T cells (Alcantara et al., 2018). However, in the presence of the antibody or protein, CAR T cells were highly enriched during culture in vitro. To investigate the phenotype of the expanded CAR T cells, the cells were washed and analyzed by flow cytometry. CD62L and CD45RA are often used to distinguish among naïve T cells (CD62Lþ CD45RAþ; TN), terminally
differentiated T cells expressing CD45RA (CD62LeCD45RAþ; TD), effector memory T cells (CD62LeCD45RAe; TEM), and central memory T cells (CD62Lþ CD45RAe; TCM) (Golubovskaya and Wu, 2016; Caccamo et al., 2018). The MM27 group increased most in the relative frequency of naïve cells (defined as CD45RA and CD62L double-positive cells) (Fig. 4A), and the frequencies of CD279 (PD1) and CD366 (Tim-3) expressing cells were reduced most (Fig. 4B).
370
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 2. Expression of CD107a and CD38 antigen on anti-CD38 CAR T cells. A: Flow cytometry was performed to assess CD107a (LAMP-1), which was a potential marker in the process of degranulation. The CD107a on self-activated anti-CD38 CAR T cells has an expression ratio of 53.69% on day 6. MOCK-Ts was used as controls. B: CD38 antigen on anti-CD38 CAR T cells was almost completely lost at the same time. C: The cell debris of anti-CD38 CAR T cells as observed by phase contrast microscopy on day 6. Scale bar: 100 mm. D: Expansion of T cells in vitro for 15 days which was transduced with truncated anti-CD38 CAR lentiviruses. **P < 0.01.
These results suggested that the anti-CD38 CAR T cells were in a naïve stage and maintained a strong capacity for differentiation. In contrast, the group cultured without the antibodies or proteins expressed relatively higher levels of PD-1 and Tim-3 than the other groups. Interestingly, anti-CD38 CAR T cells survived and expanded as a predominant CD38eCD8þ T cell subset in the group treated with additives. Furthermore, antibody or protein blockade rescued the CD4þ subpopulation within the anti-CD38 CAR T cell population, suggesting that fratricide was dependent on a skewed CD4/CD8 ratio (Fig. 4C). The relative increase in the proportion of proliferating CD4þ T cells, as well as the elimination of fratricide effect between CD4þ T and CD8þ T cells during the manufacturing of antiCD38 CAR T cells, is the most likely explanation for the difference in the ratio.
2.5. Anti-CD38 CAR T cells effectively recognize and specifically lyse CD38-positive tumor cells For cytotoxicity assessment of anti-CD38 CAR T cells, we created the A549CD38þ cell line, which is derived from A549 cells and overexpressed CD38 (Fig. 5A). The expression of the mCherry protein in A549CD38þ cells was observed by fluorescence microscopy (Fig. 5B). CAR T cells were washed several times to eliminate the effect of residual antibodies or proteins in the medium. E:T ratios of 0.5:1, 1:1, and 5:1 were established with CAR T cells and target cells, and these cells were coincubated in cytokine-free medium for 6 h, 12 h, or 24 h. In addition, reactivity was measured by monitoring proinflammatory cytokine secretion. MOCK T cells were used at the same E:T ratios and A459 cells were employed as controls. As expected, compared to the control T cells,
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
371
significantly higher proportion of degranulated anti-CD38 CAR T cells than MOCK T cells upon exposure to target cells (Fig. 5G). Furthermore, the cytotoxicity mediated by anti-CD38 CAR T cells was dose and time dependent, and the CAR T cells exhibited increased cytotoxicity as the E:T ratio increased. The CD38-negative cell line A549 was not lysed by the anti-CD38 CAR T cells, and the growth of these cells was not inhibited by the MOCK T cells (Fig. 5C). Similar to the cytokine production results, the costimulated antibody or protein group demonstrated enhanced cytotoxicity compared to their unprotected counterparts. In contrast, no cytotoxicity was detected in the experiments with the A549 cell line, which lacks CD38 expression on the cell surface (Fig. 5D). Hence, we demonstrated that anti-CD38 CAR T cells possess potent cytotoxic activity against CD38-positive tumor cells. 2.6. Effective and persistent antitumor activity of anti-CD38 CAR T cells in xenograft tumors derived from RPMI 8226 cells Anti-CD38 CAR T cells have in vivo antitumor activity, which was evaluated by inoculating NOD/SCID/g-chain/ (NPG) mice with luciferase-labeled RPMI 8226 cells (Fig. 6A). When tumors were visible, the mice were treated with anti-CD38 CAR T cells using antibody (clone MM27). MOCK T cells and NaCl were used as controls. As shown in Fig. 6B, the anti-CD38 CAR T cells induced significant regression, or even elimination, of the RPMI 8226 tumors, while tumors continued to progress in the blank and MOCK T cell groups. Intriguingly, the anti-CD38 CAR T cells appeared to modestly delay tumor growth in the groups of mice receiving the anti-CD38 CAR T cells (Fig. 6C). On day 14 after T cell infusion, the anti-CD38 CAR T cells appeared in the peripheral blood (Fig. 6D). However, after 14e28 days of treatment, most of the mice in the NaCl and MOCK T cell-treated groups had died, indicating that the survival time of tumor-bearing mice was significantly prolonged after treatment with the anti-CD38 CAR T cells (Fig. 6E). In conclusion, costimulated anti-CD38 CAR T cells, a cellular immunotherapy modality, appeared to enhance the treatment of MM in vivo. 3. Discussion Fig. 3. Strategies to avoid CAR T cell lysis and to promote cell proliferation. A: Two distinct strategies were taken to avoid fratricide which was driven by targeting and to promote anti-CD38 CAR T cell production through blocking T cell surface CD38 Ag or CAR scfv by antibodies (Ab clone MM12T or clone MM27) or proteins (H02H or H08H). The total number of cells in each group during cell proliferation and enrichment was determined. B: The transduction efficiency in each group. C: The expression of CD38 on the cell surface in each group. *P < 0.05; **P < 0.01.
the anti-CD38 CAR T cells demonstrated significant cytotoxicity against A549CD38þ cells. As shown in Fig. 5C, the MM27 group exhibited higher average cytotoxic activity against A549 CD38þ cells (41.68% ± 3.02% at 6 h and 88.35% ± 7.02% at 24 h at an E:T ratio of 5:1) than the MOCK T cell group. When tumor cells were coincubated with CAR T cells or MOCK T cells at an E:T ratio of 1:1 for 24 h, the secreted cytokines were measured by ELISA. The levels of cytokines, including IL-2 (Fig. 5D), IFN-g (Fig. 5E) and TNF-a (Fig. 5F), were significantly higher in the supernatants of cocultures containing CAR T cells and A549CD38þ cells than in those of cocultures containing MOCK T cells. The MM27 group showed the slightly stronger cytokine secretion capacity than other antibody and protein groups tested. CD107a, which is expressed on the cell surface, has been described as a marker of cytotoxic CD8þ T cell degranulation, and its expression has been shown to be strongly upregulated on the cell surface in concordance with the loss of perforin following stimulation (Lorenzo-Herrero et al., 2019). In this study, we also found a
CAR T cell therapy represents a potent and targeted immunotherapeutic approach that has been used to treat patients with hematological malignancies. The toxicity profile of CAR T cell therapy is unique but not durable. Because CD19-negative recurrence often occurs after CART-19 treatment, there are two main reasons for treatment failure with CAR T cells: relapse with antigen loss and CAR T cell population loss (Wang et al., 2017). 3.1. CD38-based fratricide decreases the expansion and enrichment of anti-CD38 CAR T cells The human CD38 Ag is prevalently expressed by T lymphocytes, plasma cells, monocytes, NK cells and other cell types. The CD38 Ag is also overexpressed in various tumor cells, including those in BNHL, MM, and some relapsed and/or refractory diseases after CAR T cell therapy to eliminate CD38-positive cells (Chini et al., 2018). This antigen is considered an attractive target for treating hematological malignancies (Bannas and Koch-Nolte, 2018). Early studies confirmed that anti-CD38 CAR T cells have a strong antitumor ability. Mihara (Mihara et al., 2009) first reported that anti-CD38 CAR T cells self-lyse by fratricide. In contrast, Drent et al. (2016) speculated that no obvious dysfunction was observed in antiCD38 CAR T cells due to the lack of expression of CD38. Another study (An et al., 2018) observed some fratricide in new nanobodybased anti-CD38 CAR T cells. In this study, fratricide of the anti-
372
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 4. A detection of subsets of CAR T cells blocked with antibodies (clone MM12T or clone MM27) or proteins (H02H or H08H) at 15 days. A: Flow cytometry was performed to assess CD62L and CD45RA simultaneously, which were used to distinguish naïve (CD62Lþ CD45RAþ; TN), effector memory (CD62LeCD45RAe; TEM), central memory (CD62Lþ CD45RAe; TCM) and terminally differentiated T cell expressing CD45RA (CD62Le CD45RAþ; TD). B: Flow cytometry was performed to assess Tim-3 and PD-1 simultaneously, which were associated with T cell exhaustion/dysfunction. C: The expression levels of CD4 and CD8 were also measured by flow cytometry.
CD38 CAR T cells occurred, and the cells were unable to proliferate effectively because of the loss of CD38 expression. The most likely reason for this loss was that the potential target-driven fratricide of the anti-CD38 CAR T cells led to a “CAR T cell - autophagic and autostimulatory/enrichment” cycle.
3.2. Antibody- or protein-mediated blockade prevents anti-CD38 CAR T cell fratricide In the current study, we tried two distinct strategies to avoid fratricide driven by targeting CD38 and to increase anti-CD38 CAR T
Fig. 5. Efficacy and anti-tumor function of anti-CD38 CAR T cells in vitro. A: A549 was overexpressed with CD38 using a lentiviral vector with an mCherry tag and sorted by flow cytometry. B: The expression of mCherry protein was detected by fluorescence microscopy to show the expression of CD38 in A549CD38þ cells (original magnification, 100). C: A549 and A549CD38þ cells were cocultured with anti-CD38 CAR T cells at effector-to-target ratios of 0.5:1, 1:1, or 5:1 for 6 h, 12 h, and 24 h. The lysis of A549CD38þ cells by anti-CD38 CAR T cells was observed at each time point. Negative effector was A459 cells, as well as target cell controls. D-F: When co-incubating the tumor cells with anti-CD38 CAR T cells or MOCK T cells at an E: T ratio of 1:1 for 24 h, the secreted cytokines were determined by ELISA, including IL-2 (D), IFN-g (E), and TNF-a (F). G: Flow cytometry was carried to detect the expression levels of CD107a in anti-CD38 CAR T cells or MOCK T cells. *P < 0.05, **P < 0.01.
374
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
Fig. 6. Strong antitumor activity of anti-CD38 CAR T cells in an MM model. A: Schematic of this study. NPG mice were inoculated intraperitoneally (i.p.) in the flank with 1 106 RPMI 8226-mCherry-luc cells on day 7. After 1 week, anti-CD38 CAR T cells treated by antibody (clone MM27) were injected via the tail vein on day 0. MOCK-Ts and NaCl were used as controls. B: Bioluminescence imaging (BLI) in each mouse was measured to study the tumor burden, and the changes of tumor burden within 6 weeks after CAR-T reinfusion were presented. C: ROI of each BLI at day 14 was analyzed. Error bars denote the s.e.m., and two-way ANOVA was chosen to compare the results. **P < 0.01. D: T cell percentage at day 14 in the peripheral blood of mice intraperitoneally injected with RPMI 8226. Unpaired t-test was used for comparison of results. **P < 0.01. E: Survival curve of mice intraperitoneally injected with RPMI 8226 cells. Results were analyzing using a Log-rank (Mantel-Cox) test. **P < 0.01.
cell production by blocking the T cell surface expression of CD38 or the CAR scFv domain with an antibody (clone MM12T or clone MM27) or a protein (H02H or H08H). The results showed that the antibodies and proteins impacted CAR T cell fratricide and
promoted CAR T cell expansion and enrichment. Furthermore, the addition of the antibody clone MM27 was most effective in improving CAR T cell yield, increasing the relative frequency of naïve cells (defined as CD62L and CD45RA double-positive cells)
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
and reducing the frequencies of CD279 (PD-1) (Catakovic et al., 2017) and CD366 (Tim-3) expressing cells (Banerjee and Kane, 2018). This result suggested that the treated anti-CD38 CAR T cells had improved capacities for self-renewal and long-term survival. These data have powerful implications on the large-scale expansion of CAR T cells after tumor cell recognition. In addition, both protein treatments generated comparable anti-CD38 CAR T cell populations. Furthermore, these methods have been used in clinical trials to efficiently and reproducibly produce anti-CD38 CAR T cells. Previous preclinical studies have indicated that differences in CAR T efficacy depend on the different populations of CD4þ T cells and CD8þ T cells. The manufacture of CAR T cell products with a defined 1:1 ratio of CD4þ/CD8þ T cells was demonstrated to be the most efficacious approach (Sommermeyer et al., 2016). In this study, CD4þ anti-CD38 CAR T cells suffered severe lysis in the absence of antibodies or proteins protection during culture. In addition, a subset of CD8þ CAR T cells expanded and proliferated. The lack of helper CD4þ T cells might reduce the antitumor ability and persistence of generated CAR T cells. Conversely, CD4þ CAR T cells were rescued by pretreatment with antibodies or proteins during CAR T cell culture. These results indicate that fratricide is dependent on the CD4þ/CD8þ T cell ratio. Differences in this ratio are most likely due to the proliferation of CD4þ T cells or the reduction in CD4þ T cell numbers caused by CD8þ T cells. 3.3. Specific cytotoxicity of anti-CD38 CAR T cells against CD38þ cell lines In the current study, we constructed anti-CD38 CAR T cells and evaluated their antitumor activity and persistence. The anti-CD38 CAR T cells were cocultured with CD38þ tumor cell lines to assay cytolysis. It has consistently been demonstrated by in vitro results that CD38þ cells can be lysed efficiently and specifically by antiCD38 CAR T cells in a dose- and time-dependent manner. Similarly, the antitumor effect of CAR T cells on primary CD38þ tumor cells has also been demonstrated. The antitumor effect of CAR T cells is positively correlated with the secretion levels of various cytokines, and the levels of the proinflammatory factors IL-2, IFN-g and TNF-a were significantly upregulated in the supernatants of A549CD38þ cells. Cell surface-expressed CD107a is considered to be a marker of CD8þ T cell degranulation. In this experiment, we found a significantly higher proportion of degranulated anti-CD38 CAR T cells than degranulated MOCK T cells upon exposure to target cells. Similar antitumor activity was observed in a NOD/SCID/g-chain/ (NPG) mouse xenograft model. Specificity was confirmed by using MOCK T cells and A549 cells as negative controls. In conclusion, targeting CD38 with CAR T cells is an enticing and novel approach to further improve antitumor effects. However, the target-driven fratricide of anti-CD38 CAR T cells remains a major problem. Our strategy avoided cell lysis and obtained a sufficient preparation of cells for CAR T cell manufacturing, especially in the presence of the antibody clone MM27. We will verify this strategy in future experiments. Anti-CD38 CAR T cells still face significant challenges in future clinical studies. Since the CD38 Ag is widely expressed in normal cells and tissues, there is a risk of infused CAR T cells attacking these cells and tissues. Most importantly, in myeloid progenitors, CD38 is a common differentiation antigen (Miyawaki et al., 2017), and abnormal hematopoiesis may result after targeting the CD38 Ag. In this study, immunotherapy mediated by anti-CD38 CAR T cells has the potential to eliminate CD38þ tumor cells and overcome target-driven fratricide of CAR T cells through blockade of T cell surface-expressed CD38 Ag or CAR scFv by antibodies or proteins. Anti-CD38 CAR T cells demonstrated specific and strong antitumor
375
effects in vitro and in vivo. The results will help to further enhance the expansion, persistence and function of anti-CD38 CAR T cells in subsequent clinical trials (NCT03754764). 4. Methods and materials 4.1. Cell lines and culture reagents The human MM cell line RPMI 8226 (CD38-positive) was used in immune-based assays. MM cells were cultured in RPMI medium (Sigma-Aldrich, St. Louis, MO, USA) containing 10% fetal calf serum (Atlas Biologicals, USA). The human lung adenocarcinoma cell line A549 was used as the antigen-negative control. CD38overexpressing A549 cells were constructed with a lentiviral vector with an mCherry tag and named A549CD38þ cells. The immortalized normal fetal renal 293T cell line was cultured in DMEM for lentiviral packaging. CD38-positive primary tumor cells were obtained from a patient. Complete medium was used for the culture of all cell lines: 100 mg/mL streptomycin sulfate, 100 U/mL penicillin and 10% heat inactivated fetal bovine serum (FBS) were added to supplemented RPMI-1640 medium. CD38 proteins (H02H and H08H) and antibodies (clone MM12T and MM27) were purchased from Beijing Sino Biological Co., Ltd. 4.2. Constructing anti-CD38 CAR T cells and producing lentiviruses The anti-CD38 CAR consisted of the human anti-CD38 scFv followed by the intracellular signaling motifs of human CD137 and CD3z (Fig. 2A), which have been described previously (Wang et al., 2015; Zhang et al., 2016). The pWPT-anti-CD38 CAR plasmid and ps-PAX2 packaging plasmid were transfected into 293 T cells. All the lentiviruses in this study were obtained from concentrated stocks. Before experimentation or titration, viruses were aliquoted and stored at 80 C. 4.3. Transduction of human T cells Primary human T cells were isolated from healthy normal donors. Peripheral blood mononuclear cells (PBMCs) were suspended in XeVIVO 15 medium directly (Lonza, USA) with an anti-CD3 monoclonal antibody (mAb; 500 ng/mL) and recombinant human IL-2 (300 U/mL) (PeproTech, USA). Transduction mediated by a lentivirus was performed on day 0. Briefly, in the presence of protamine sulfate (Sigma-Aldrich) at a concentration of 10 mg/mL, 1 106 T cells were infected with lentiviral vectors encoding CAR constructs overnight. After transduction, the cultures were transferred to fresh XeVIVO 15 medium supplemented with IL-2 addition every other day, and the cell density was maintained at 0.5e1 106 cells/mL. 4.4. Antibodies or proteins inhibition assays Two distinct strategies were used to avoid target-driven fratricide. The first strategy was to use the mouse IgG1-type antibody clones MM12T or MM27 to block the anti-CD38 CAR on the T cell surface. The second strategy was to use the protein H02H or H08H to block the cell-transduced scFv domain of the CAR. In the process of cell culture, the appropriate protein or antibody was added to the CAR T cell culture medium at the end of cell transduction. MOCK T cell and no additive CAR T cell groups were established at the same time. In coculture experiments, to eliminate the effects of residual antibody or protein, CAR T cells were washed several times in medium.
376
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377
4.5. Flow cytometric analysis The following fluorochrome-conjugated antibodies purchased from BD (Biosciences, USA) were used in this study: APC-Cy7 antihuman CD3, FITC anti-human CD4, PE anti-human CD8, PE antihuman CD366 (Tim-3). APC anti-human CD269 (PD-1), APC antihuman CD45RA, PE anti-human CD62L, and APC anti-human CD107a. Annexin V-PE and 7-Aminoactinomycin D (7-AAD) were used for viability assessment (Crowley et al., 2016). In experiments with the addition of the mouse IgG1-type antibody clones MM12T and MM27, a goat anti-mouse IgG1 antibody (FITC) was chosen to analyze CD38 expression on cells. In the remaining groups, including the MOCK T cells with the addition of protein H02H or H08H groups and the no additive CAR T cell groups, the CAR T cells were stained with a PE-conjugated anti-human CD38 antibody. A recombinant CD38-Fc protein and a goat anti-mouse Fc (APC) antibody were used to evaluate surface expression. All analyses used matched secondary and isotype antibodies. Biotin-spaffinipure goat anti-mouse IgG F(ab')2 fragment and a PEsreptavidin antibody were used to detect MOCK T cells in the following experiments. A Beckman Coulter CytoFlex flow cytometer was used to perform flow cytometry, and FlowJo software was used to analyze the data (Version 10. 0. 7, FlowJo, Ashland, OR, USA). 4.6. Cytokine release assays Anti-CD38 CAR T cells and target cells were cocultured in a 1:1 ratio, followed by evaluation with a cytokine release assay. The cells were cultured in 96-well plates, with 200 mL of RPMI-1640 medium in each well. When the experiment ended, an ELISA (BioLegend, USA) was performed using cell-free coculture supernatants to measure the secretion of TNF-a, IL-2 and IFN-g. The results were measured three times and were expressed as an average. 4.7. Cytotoxicity assays Adherent A549 and A549CD38þ cells in each of the above mentioned groups were cultured in a flat-bottomed 96-well plate for 20 h at E:T ratios of 0.5:1, 1:1, and 5:1 for 6 h, 12 h, or 24 h. AntiCD38 CAR T cells were washed, and the remaining adherent A549CD38þ cells were labeled with alamar Blue (ThermoFisher Scientific, USA) in serum-free medium (Xu et al., 2015; Breman et al., 2018) for 4 h. The Infinite M200 Pro (TECAN, Switzerland) was used to measure viable cells at 530 nm, and then the relative cytolytic activity was calculated. 4.8. Xenograft model of MM To establish a xenotransplanted tumor model, female NPG mice (6e8 weeks old) were obtained from Beijing Vitalstar Biotechnology Co., Ltd. and inoculated intraperitoneally (i.p.) in the flank with 1 106 RPMI 8226-mCherry-luc cells on day 7. The tumors became palpable at approximately 1 week, and the mice were injected via the tail vein with anti-CD38 CAR T cells or MOCK T cells (1 106 cells suspended in 100 mL of PBS) on day 0. Bioluminescence imaging (BLI) was used to monitor tumor growth. We performed the animal experiments according to the Animal Protection Guidelines of the People's Liberation Army General Hospital (PLAGH). 4.9. Statistical analysis All experiments were performed three times, and the results are presented as the mean ± SD. Differences among groups were examined using one-way analysis of variance (ANOVA). Differences
in the number of transferred T cells were evaluated by Student's ttest. Analyses were performed with IBM SPSS 23 software (IBM, USA), and a two-tailed P < 0. 01 was considered statistically significant. Acknowledgments We thank all the patients for consenting to provide tissue samples. This work was supported by the grants from the National Natural Science Foundation of China (Nos. 81830002 and 31870873 to W.D.H.) and the National Key Research and Development Program of China (Nos. 2016YFC1303501 and 2016YFC1303504 to WDH, and No. 2017YFC0909803 to W.Y.). References Alcantara, M., Tesio, M., June, C.H., Houot, R., 2018. CAR T-cells for T-cell malignancies: challenges in distinguishing between therapeutic, normal, and neoplastic T-cells. Leukemia 32, 2307e2315. An, N., Hou, Y.N., Zhang, Q.X., Li, T., Zhang, Q.L., Fang, C., Chen, H., Lee, H.C., Zhao, Y.J., Du, X., 2018. Anti-multiple myeloma activity of nanobody-based anti-CD38 chimeric antigen receptor T cells. Mol. Pharm. 15, 4577e4588. Atanackovic, D., Steinbach, M., Radhakrishnan, S.V., Luetkens, T., 2016. Immunotherapies targeting CD38 in multiple myeloma. OncoImmunology 5 e1217374. Banerjee, H., Kane, L.P., 2018. Immune Regulation by Tim-3. F1000Res, vol. 7, p. 316. Bannas, P., Koch-Nolte, F., 2018. Perspectives for the development of CD38-Specific heavy chain antibodies as therapeutics for multiple myeloma. Front. Immunol. 9, 2559. Blatt, K., Menzl, I., Eisenwort, G., Cerny-Reiterer, S., Herrmann, H., Herndlhofer, S., Stefanzl, G., Sadovnik, I., Berger, D., Keller, A., Hauswirth, A., Hoermann, G., Willmann, M., Rulicke, T., Sill, H., Sperr, W.R., Mannhalter, C., Melo, J.V., Jager, U., Sexl, V., Valent, P., 2018. Phenotyping and target expression profiling of CD34þ/ CD38- and CD34þ/CD38þ stem- and progenitor cells in acute lymphoblastic leukemia. Neoplasia 20, 632e642. Bonello, F., D'Agostino, M., Moscvin, M., Cerrato, C., Boccadoro, M., Gay, F., 2018. CD38 as an immunotherapeutic target in multiple myeloma. Expert Opin. Biol. Ther. 18, 1209e1221. Breman, E., Demoulin, B., Agaugue, S., Mauen, S., Michaux, A., Springuel, L., Houssa, J., Huberty, F., Jacques-Hespel, C., Marchand, C., Marijsse, J., Nguyen, T., Ramelot, N., Violle, B., Daro, D., De Waele, P., Gilham, D.E., Steenwinckel, V., 2018. Overcoming target driven fratricide for T cell therapy. Front. Immunol. 9, 2940. Brudno, J.N., Kochenderfer, J.N., 2019. Recent advances in CAR T-cell toxicity: mechanisms, manifestations and management. Blood Rev. 34, 45e55. Caccamo, N., Joosten, S.A., Ottenhoff, T.H.M., Dieli, F., 2018. Atypical human effector/ memory CD4þ T cells with a naive-like phenotype. Front. Immunol. 9, 2832. Catakovic, K., Klieser, E., Neureiter, D., Geisberger, R., 2017. T cell exhaustion: from pathophysiological basics to tumor immunotherapy. Cell Commun. Signal. 15, 1. Chehab, S., Panjic, E.H., Gleason, C., Lonial, S., Nooka, A.K., 2018. Daratumumab and its use in the treatment of relapsed and/or refractory multiple myeloma. Future Oncol. 14, 3111e3121. Chini, E.N., Chini, C.C.S., Espindola Netto, J.M., de Oliveira, G.C., van Schooten, W., 2018. The pharmacology of CD38/NADase: an emerging target in cancer and diseases of aging. Trends Pharmacol. Sci. 39, 424e436. Crowley, L.C., Marfell, B.J., Scott, A.P., Waterhouse, N.J., 2016. Quantitation of apoptosis and necrosis by annexin V binding, propidium iodide uptake, and flow cytometry. Cold Spring Harb. Protoc. 2016 https://doi.org/10.1101/pdb. prot087288. Drent, E., Groen, R.W., Noort, W.A., Themeli, M., Lammerts van Bueren, J.J., Parren, P.W., Kuball, J., Sebestyen, Z., Yuan, H., de Bruijn, J., van de Donk, N.W., Martens, A.C., Lokhorst, H.M., Mutis, T., 2016. Pre-clinical evaluation of CD38 chimeric antigen receptor engineered T cells for the treatment of multiple myeloma. Haematologica 101, 616e625. Feng, K.C., Guo, Y.L., Liu, Y., Dai, H.R., Wang, Y., Lv, H.Y., Huang, J.H., Yang, Q.M., Han, W.D., 2017. Cocktail treatment with EGFR-specific and CD133-specific chimeric antigen receptor-modified T cells in a patient with advanced cholangiocarcinoma. J. Hematol. Oncol. 10, 4. Golubovskaya, V., Wu, L., 2016. Different subsets of T cells, memory, effector functions, and CAR-T immunotherapy. Cancers (Basel) 8, 36. Han, X., Wang, Y., Han, W.D., 2018. Chimeric antigen receptor modified T-cells for cancer treatment. Chronic Dis. Transl. Med. 4, 225e243. Kloss, S., Oberschmidt, O., Dahlke, J., Vu, X.K., Neudoerfl, C., Kloos, A., Gardlowski, T., Matthies, N., Heuser, M., Meyer, J., Sauer, M., Falk, C., Koehl, U., Schambach, A., Morgan, M.A., 2019. Preclinical assessment of suitable natural killer (NK) cell sources for chimeric antigen receptor (CAR)-NK-based "off-the-shelf" AML immunotherapies. Hum. Gene Ther. 30. https://doi.org/10.1089/hum.2018.247. Lorenzo-Herrero, S., Sordo-Bahamonde, C., Gonzalez, S., Lopez-Soto, A., 2019. CD107a degranulation assay to evaluate immune cell antitumor activity. Methods Mol. Biol. 1884, 119e130. McHayleh, W., Bedi, P., Sehgal, R., Solh, M., 2019. Chimeric antigen receptor T-cells:
Z. Gao et al. / Journal of Genetics and Genomics 46 (2019) 367e377 the future is now. J. Clin. Med. 8 pii: E207. Mele, S., Devereux, S., Pepper, A.G., Infante, E., Ridley, A.J., 2018. Calcium-RasGRP2Rap1 signaling mediates CD38-induced migration of chronic lymphocytic leukemia cells. Blood Adv. 2, 1551e1561. Mihara, K., Bhattacharyya, J., Kitanaka, A., Yanagihara, K., Kubo, T., Takei, Y., Asaoku, H., Takihara, Y., Kimura, A., 2012. T-cell immunotherapy with a chimeric receptor against CD38 is effective in eliminating myeloma cells. Leukemia 26, 365e367. Mihara, K., Yanagihara, K., Takigahira, M., Imai, C., Kitanaka, A., Takihara, Y., Kimura, A., 2009. Activated T-cell-mediated immunotherapy with a chimeric receptor against CD38 in B-cell non-Hodgkin lymphoma. J. Immunother. 32, 737e743. Mihara, K., Yanagihara, K., Takigahira, M., Kitanaka, A., Imai, C., Bhattacharyya, J., Kubo, T., Takei, Y., Yasunaga, S., Takihara, Y., Kimura, A., 2010. Synergistic and persistent effect of T-cell immunotherapy with anti-CD19 or anti-CD38 chimeric receptor in conjunction with rituximab on B-cell non-Hodgkin lymphoma. Br. J. Haematol. 151, 37e46. Miyawaki, K., Iwasaki, H., Jiromaru, T., Kusumoto, H., Yurino, A., Sugio, T., Uehara, Y., Odawara, J., Daitoku, S., Kunisaki, Y., Mori, Y., Arinobu, Y., Tsuzuki, H., Kikushige, Y., Iino, T., Kato, K., Takenaka, K., Miyamoto, T., Maeda, T., Akashi, K., 2017. Identification of unipotent megakaryocyte progenitors in human hematopoiesis. Blood 129, 3332e3343. Morandi, F., Horenstein, A.L., Costa, F., Giuliani, N., Pistoia, V., Malavasi, F., 2018. CD38: a target for immunotherapeutic approaches in multiple myeloma. Front. Immunol. 9, 2722. Naik, J., Themeli, M., de Jong-Korlaar, R., Ruiter, R.W.J., Poddighe, P.J., Yuan, H., de Bruijn, J.D., Ossenkoppele, G.J., Zweegman, S., Smit, L., Mutis, T., Martens, A.C.M., van de Donk, N., Groen, R.W.J., 2018. CD38 as a therapeutic target for adult acute myeloid leukemia and T-cell acute lymphoblastic leukemia. Haematologica 104, e100ee103. Pick, M., Vainstein, V., Goldschmidt, N., Lavie, D., Libster, D., Gural, A., Grisariu, S., Avni, B., Ben Yehuda, D., Gatt, M.E., 2018. Daratumumab resistance is frequent in advanced-stage multiple myeloma patients irrespective of CD38 expression and is related to dismal prognosis. Eur. J. Haematol. 100, 494e501.
377
Plesner, T., Krejcik, J., 2018. Daratumumab for the treatment of multiple myeloma. Front. Immunol. 9, 1228. Roddie, C., O'Reilly, M., Dias Alves Pinto, J., Vispute, K., Lowdell, M., 2019. Manufacturing chimeric antigen receptor T cells: issues and challenges. Cytotherapy 21, 327e340. Sommermeyer, D., Hudecek, M., Kosasih, P.L., Gogishvili, T., Maloney, D.G., Turtle, C.J., Riddell, S.R., 2016. Chimeric antigen receptor-modified T cells derived from defined CD8þ and CD4þ subsets confer superior antitumor reactivity in vivo. Leukemia 30, 492e500. van de Donk, N., 2018. Immunomodulatory effects of CD38-targeting antibodies. Immunol. Lett. 199, 16e22. van de Donk, N., Usmani, S.Z., 2018. CD38 antibodies in multiple myeloma: mechanisms of action and modes of resistance. Front. Immunol. 9, 2134. Wang, J., Hu, Y., Huang, H., 2017. Acute lymphoblastic leukemia relapse after CD19targeted chimeric antigen receptor T cell therapy. J. Leukoc. Biol. 102, 1347e1356. Wang, Q.S., Wang, Y., Lv, H.Y., Han, Q.W., Fan, H., Guo, B., Wang, L.L., Han, W.D., 2015. Treatment of CD33-directed chimeric antigen receptor-modified T cells in one patient with relapsed and refractory acute myeloid leukemia. Mol. Ther. 23, 184e191. Xu, M., McCanna, D.J., Sivak, J.G., 2015. Use of the viability reagent prestoplue in comparison with alamarblue and MTT to assess the viability of human corneal epithelial cells. J. Pharmacol. Toxicol. Methods 71, 1e7. Zaja, F., Tabanelli, V., Agostinelli, C., Calleri, A., Chiappella, A., Varettoni, M., Luigi Zinzani, P., Volpetti, S., Sabattini, E., Fanin, R., Pileri, S.A., 2017. CD38, BCL-2, PD1, and PD-1L expression in nodal peripheral T-cell lymphoma: possible biomarkers for novel targeted therapies? Am. J. Hematol. 92, E1eE2. Zhang, C., Kasi, A., 2018. Chimeric Antigen Receptor (CAR) T-Cell Therapy. StatPearls, Treasure Island (FL). Zhang, W.Y., Wang, Y., Guo, Y.L., Dai, H.R., Yang, Q.M., Zhang, Y.J., Zhang, Y., Chen, M.X., Wang, C.M., Feng, K.C., Li, S.X., Liu, Y., Shi, F.X., Luo, C., Han, W.D., 2016. Treatment of CD20-directed chimeric antigen receptor-modified T cells in patients with relapsed or refractory B-cell non-Hodgkin lymphoma: an early phase IIa trial report. Signal Transduct. Target. Ther. 1, 16002.