Rapid generation of clinical-grade antiviral T cells: selection of suitable T-cell donors and GMP-compliant manufacturing of antiviral T cells

Abstract

Background: The adoptive transfer of allogeneic antiviral T lymphocytes derived from seropositive donors can safely and effectively reduce or prevent the clinical manifestation of viral infections or reactivations in immunocompromised recipients after hematopoietic stem cell (HSCT) or solid organ transplantation (SOT). Allogeneic third party T-cell donors offer an alternative option for patients receiving an allogeneic cord blood transplant or a transplant from a virus-seronegative donor and since donor blood is generally not available for solid organ recipients. Therefore we established a registry of potential third-party T-cell donors (allogeneic cell registry, alloCELL) providing detailed data on the assessment of a specific individual memory T-cell repertoire in response to antigens of cytomegalovirus (CMV), Epstein-Barr virus (EBV), adenovirus (ADV), and human herpesvirus (HHV) 6.

Methods: To obtain a manufacturing license according to the German Medicinal Products Act, the enrichment of clinical-grade CMV-specific T cells from three healthy CMV-seropositive donors was performed aseptically under GMP conditions using the CliniMACS cytokine capture system (CCS) after restimulation with an overlapping peptide pool of the immunodominant CMVpp65 antigen. Potential T-cell donors were selected from alloCELL and defined as eligible for clinical-grade antiviral T-cell generation if the peripheral fraction of IFN-γ(+) T cells exceeded 0.03% of CD3(+) lymphocytes as determined by IFN-γ cytokine secretion assay.

Results: Starting with low concentration of IFN-γ(+) T cells (0.07-1.11%) we achieved 81.2%, 19.2%, and 63.1% IFN-γ(+)CD3(+) T cells (1.42 × 10(6), 0.05 × 10(6), and 1.15 × 10(6)) after enrichment. Using the CMVpp65 peptide pool for restimulation resulted in the activation of more CMV-specific CD8(+) than CD4(+) memory T cells, both of which were effectively enriched to a total of 81.0% CD8(+)IFN-γ(+) and 38.4% CD4(+)IFN-γ(+) T cells. In addition to T cells and NKT cells, all preparations contained acceptably low percentages of contaminating B cells, granulocytes, monocytes, and NK cells. The enriched T-cell products were stable over 72 h with respect to viability and ratio of T lymphocytes.

Conclusions: The generation of antiviral CD4(+) and CD8(+) T cells by CliniMACS CCS can be extended to a broad spectrum of common pathogen-derived peptide pools in single or multiple applications to facilitate and enhance the efficacy of adoptive T-cell immunotherapy.

Figure 1

Protocol for the rapid manufacture of clinical-grade antigen-specific T cells. A three-step protocol for the rapid generation of clinical-grade antiviral T cells was established to facilitate the manufacture of specific T cells for adoptive transfer in pre-monitored patients. First Step: Selection of potential T-cell donors from the alloCELL registry (HLA type, virus serology and virus-specific T-cell response). Second Step: Verification of the donor’s specific T-cell frequencies (donor from alloCELL, stem cell or family donor) and prediction of the donor’s T-cell enrichment efficiency by small-scale MiniMACS CSA. A T-cell donor is classified as eligible if (a) the peripheral frequency of virus-specific IFN-γ⁺ T cells ≥0.03% of total CD3⁺ T cells and (b) the restimulation efficiency is twice as much as the unstimulated control. Third Step: Manufacturing of clinical-grade antiviral T cells by large-scale CliniMACS CCS. A CliniMACS CCS-enriched T-cell fraction (TCF) is classified as eligible if (a) number of viable IFN-γ⁺ T cells >1 × 10⁴ and (b) the number of viable IFN-γ⁻ T cells ≤2 × 10⁷.

Figure 2

Gating strategy established for flow cytometric quality and in-process control regarding the CliniMACS CCS validation. Samples of the collected CliniMACS CCS fraction were analysed by flow cytometry using the Quality control panel QCP-A/A⁻ and the represented gating strategy. All cell fractions (leukapheresis, original fraction (OF), T-cell fraction (TCF), negative fraction (NF), waste fraction (WF), 48 h, 54 h, and 72 h post-leukapheresis (Stabi48, Stabi54, and Stabi72)) were stained with specific antibodies to visualize IFN-γ⁺ T cells. In the first plot, cells were analysed by 7AAD viability staining to determine the live versus dead cells, followed by gating cells based upon CD45 expression to identify CD45⁺ leukocytes in the total viable 7AAD⁻ population. In the next gating step, T cells were selected based on CD3 expression. CD3⁺CD56⁺ NKT cells were gated out using a dump channel. CD4 and CD8 surface expression was then determined from this gated population. IFN-γ⁺ T cells were gated on CD3⁺CD56⁻ T cells and on the CD4⁺ and CD8⁺ subpopulation of CD3⁺CD56⁻ T cells. The axes of the dot plots are biexponential.

Figure 3

Flow cytometric quality and in-process control of IFN-γ-based CliniMACS CCS enrichment of CMV-specific T cells. IFN-γ⁺ CMV-specific T cells were isolated from leukapheresis by large-scale GMP-grade CliniMACS CCS- and small-scale MiniMACS CSA-based process. Flow cytometric analysis was performed with all CliniMACS CCS and MiniMACS CSA fractions (leukapheresis, original fraction (OF), T-cell fraction (TCF), negative fraction (NF), waste fraction (WF), 48 h, 54 h, and 72 h post-leukapheresis (Stabi48, Stabi54, and Stabi72)) by using the quality control panel (QCP) -A, QCP-B and QCP-C-. The results of the representative analysis of the leukapheresis and TCF by using the QCP-A panel are shown (n = 3). As a control the QCP-A⁻ was used as fluorescence minus one (FMO) for IFN-γ. Dot plots show the qualitative analysis of IFN-γ-secreting CMV-specific T cells [%]. CD3⁺IFN-γ⁺ percentages were defined on viable CD3⁺ T cells, and CD8⁺IFN-γ⁺ and CD4⁺IFN-γ⁺ percentages were defined on viable CD4⁺ and viable CD8⁺ T-cells, respectively. IFN-γ⁻secreting T cells are shown in the gate represented on each dot plot.

Figure 4

Efficiency and outcomes of CliniMACS CCS validation for CMVpp65-specific T-cell enrichment. The percentage of IFN-γ secreting CMVpp65-specific T cells was detected after four hours of ex vivo stimulation with CMVpp65_pp using the QCP-A/A^- panel. The IFN-γ-based CliniMACS CCS and MiniMACS CSA systems were used for the isolation of IFN-γ-secreting CMVpp65-specific T cells. (A) The numbers of IFN-γ⁺ cells [x10⁶] within the CD3, CD4 and CD8 T-cell populations were analysed in all collected CliniMACS CCS fractions (leukapheresis, original fraction (OF), T-cell fraction (TCF), negative fraction (NF), waste fraction (WF), 48 h, 54 h, and 72 h post-leukapheresis (Stabi48, Stabi54, Stabi72)) to determine the efficiency of the CliniMACS process. Data sets of the representative analysis of the leukapheresis, OF, TCF, NF, WF fractions are shown (B) Outcome of IFN-γ-based CliniMACS CCS and MiniMACS CSA processes regarding the percentage [%] of IFN-γ⁺ cells among the CD3, CD4 and CD8 T-cell populations in samples collected at different steps of the validation process (leukapheresis, OF, TCF, NF, WF, Stabi48, Stabi54, Stabi72). Data sets of the representative analysis of the leukapheresis and TCF are shown. The results of independent experiments (n = 3) are expressed as the mean frequency of IFN-γ⁺ T cells ± SD. Asterisks indicate statistically significant differences between samples before and after T-cell enrichment (*p < 0.05).

Figure 5

Analysis of product stability. Stability of the TCF was analysed after 48 h, 54 h and 72 h of the start of leukapheresis with respect to product viability [%], frequency of CD3⁺CD56⁻ T cells [%] in CD45⁺ leukocytes and IFN-γ^+/− T cells [%] in CD3⁺CD56⁻ lymphocytes. The results of independent experiments are expressed as the mean frequency [%] of viability, T cells and IFN-γ^+/− T cells with regard to the different time points of storage.

Figure 6

Post-processing assessment of leukocyte subsets in the TCF. Fractions collected during the CliniMACS CCS process (leukapheresis, original fraction (OF), T-cell fraction (TCF), negative fraction (NF), waste fraction (WF), 48 h, 54 h, and 72 h post-leukapheresis (Stabi48, Stabi54, Stabi72)) were assessed for leukocyte subsets including: CD3⁺ T cells, CD3⁺CD56⁺ NKT cells, CD3⁻CD56⁺ NK cells, CD19⁺ B cells, CD33⁺ granulocytes and CD14⁺ monocytes. (A) The compositions of the leukocyte subsets in the Leukapheresis products and the TCFs and (B) the log depletion in cell numbers of leukocyte subsets after CliniMACS CCS enrichment are shown. The results of independent experiments are expressed as the number [x10⁶] of IFN-γ⁺ T cells ± SD and the fold decrease [log] of cell numbers in leukocyte subsets. Asterisks indicate statistically significant differences between T cells and other leukocyte subsets (*p < 0.05).