Direct Isolation of Donor-Derived Antigen - Specific T Cells and Their Adoptive Transfer for Treatment or Prophylaxis of CMV Infection Following Allogeneic Transplantation.

Reactivation of CMV is a significant cause of morbidity and mortality following allogeneic hematopoietic stem cell transplantation. Available antiviral drug therapy is effective but toxic, and contributes to the negative impact of CMV infection in affected patients. Virus-specific T lymphocytes are necessary for the control of viral reactivation, and therapeutic strategies using adoptive transfer of donor derived CMV-specific T cells are being explored. Most methods to produce donor-derived antigen-specific T cells involve several weeks of in vitro culture. The current method involves a 20hr incubation of donor-derived PBMCs with CMV-pp65 protein with subsequent isolation of T cells secreting interferon-gamma (IFNγ) by CliniMACS using IFNγ capture microbeads (Miltenyi Biotec). This technique permits rapid isolation of an enriched IFNγ secreting T cell product, manufactured to clinical grade, which is then cryopreserved in dosed aliquots for subsequent infusion. A single arm phase I study is currently underway, with CMV-specific T cells given pre-emptively at first detection (by quantitative PCR) of CMV DNA in peripheral blood, or at day +40–50 as prophylaxis. Cells are infused at an initial dose of 1x10⁴ CD3⁺/kg recipient weight, and CMV monitored by weekly PCR. Antiviral drug therapy is instituted if the viral load subsequently rises above threshold, according to institutional guidelines. Patients have received at least one dose of CMV-T cells and all are alive and well. Follow-up data are available currently on 7. These 7 patients had all received T cell deplete regimens carrying a CMV reactivation risk of >85%. The mean yield of cells following enrichment was 15.6 x10⁶ (range 5.2–26.7) of which 82.3% were CD3⁺, 22.0% were CD4⁺/IFNγ⁺ and 10.6% were CD8⁺/IFNγ⁺, giving a total mean CMV-reactive cell yield of 3.4 x10⁶ per donor. Following infusion, in vivo expansion of CMV-reactive T cells was observed in all patients. CMV-reactive cells represented a mean of 9.0% of CD4⁺ cells and 7.3% of CD8⁺ cells by 2–4 weeks post-infusion; the result of in vivo expansions of CMV-reactive cells of 700- to 5000-fold by week 4 post infusion. Two patients had CMV-specific cells infused prophylactically at day 40–50 and expansions of CMV-reactive T cells were seen in both despite consistent absence of detectable viral DNA in peripheral blood. Of the remaining 5 patients, all received antiviral therapy for the initial reactivation, although in 4 patients this was required for a significantly shorter period than in historical controls (11–14 days). In the final patient, more prolonged treatment was required (33 days) and a second reactivation also required drug therapy- in this patient, the expansion of CMV reactive T cells was substantially delayed compared to the other 6. Two other patients had a further CMV reactivation event. One followed prednisolone therapy for acute grade II graft-versus-host disease (in keeping with expected rates of acute GVHD following the primary allograft procedure) and required antiviral drug treatment. The third patient received no treatment and cleared virus following a further in vivo expansion of CMV-reactive T cells, suggesting the presence of functional memory cells. CMV-reactive T cells have been detectable up to the final 6 month sampling date. This technique provides a rapid source of clinical-grade CMV-reactive CD4⁺ and CD8⁺ T cells which early evidence suggests provide effective antiviral immunity.