Infection remains a major challenge after hematopoietic stem cell transplantation . In transplant recipients, reactivation of herpes viruses, especially of the human cytomegalovirus (CMV), is associated with severe and fatal complications. CMV is a life-long persisting virus with a seroprevalence between 45 and 100% in the general adult population . Whereas symptomatic CMV infection of healthy individuals is rare, in immunodeficient patients, such as transplant recipients, CMV infection or reactivation commonly manifests as a life-threatening disease affecting different organ systems. Transplant recipients with CMV viremia are usually treated with antiviral drugs . However, myelotoxicity as well as mutations of the viral DNA sequence may limit the use of these drugs [3,4,5]. Thus, in selected cases alternative therapeutic options are mandatory.
We here present the follow-up of a 21-year-old female with acute myeloid leukemia, who was treated with CMV-specific T cells (virus-specific T cells, VSTs). Written informed patient consent has been obtained. Viral load and cellular CMV immunity were closely monitored for almost 3 years. The patient was transplanted in cytomorphologic complete remission but with cytogenetically minimal residual disease. She received allogeneic peripheral blood stem cells from a 9/10 HLA-matched, unrelated male donor in May 2015. Myeloablative conditioning consisted of fludarabin, carmustine, antithymocyte globulin, and melphalan, and the immunosuppressive regimen included cyclosporine A and budesonide. Anti-infective prophylaxis/therapy included amoxicillin, metronidazole, itraconazole, valganciclovir, cotrimoxazol, calciumfolinate, and pentamidine. She received 9.4 × 106 CD34+ cells/kg body weight. Three weeks after peripheral blood stem cell transplantation (PBSCT) the patient suffered from acute graft vs. host disease (GvHD) of the skin (stage 1, grade 1), which did not require systemic treatment. The unrelated donor was CMV IgM- and IgG-negative, and the patient was CMV IgM-negative but IgG-positive. Thus, the recipient was at high risk of CMV reactivation. In July 2015 anti-CMV IgM and IgG became positive in the recipient and the viral load reached up to 1,022,908 copies (1,595,800 IU)/ml (day 35) as shown in Fig. 1. Despite antiviral therapy (sequentially with valganciclovir, ganciclovir, and foscarnet) and CMV-specific immunoglobulins no adequate decrease of the viral load was observed. At the peak of CMV reactivation, the patient suffered from a generalized seizure, which was considered cyclosporine-related. Treatment with cyclosporine and budesonide was switched to mycophenolate mofetil and prednisone. The patient received levetirazepam and no further seizure occurred.
It was then decided to treat the patient with CMV-specific lymphocytes of her HLA haploidentical sister who was CMV IgG-positive. Manufacturing of CMV-specific T cells was carried out at Hannover Medical School with the CliniMACS® Plus Instrument and GMP PepTivator® HCMVpp65 for antigenic restimulation, as described previously [6, 7]. Enrichment of IFN-γ-secreting CMV-specific T cells was performed by immunomagnetic separation using the CliniMACS Cytokine Capture System (Miltenyi Biotec, Bergisch Gladbach, Germany), consisting of the CliniMACS Catchmatrix and IFN-γ Enrichment Reagents. The final T-cell product had a viability of 80% with a purity of 41.8% CMV-specific IFN-γ+ T cells. Selection of the donor was based on the CMV serostatus and the presence of CMV-specific T cells as specified in Table 1. The patient received 2.5 × 104 CD3+ cells/kg body weight (12.3 ml) of CMV-VSTs on day 96 after PBSCT. CMV DNA, cellular CMV immunity, and leukocyte numbers were monitored before and after transfer of CMV-specific T cells. To determine viral load, CMV DNA was purified from blood samples using the Abbott m2000sp automated nucleic acid extraction system (Abbott, Wiesbaden, Germany). The viral load was quantified with the Abbott m2000rt real-time PCR system using the Abbott RealTime CMV amplification reagent kit according to the manufacturer’s instructions. The detection limit of this assay is 40 copies (62.4 IU)/ml.
After infusion of VSTs CMV viral load decreased until day 137 after transplantation (day 41 after VSTs) and reached a minimum of 63 copies (98 IU)/ml (Fig. 1a). On day 141 after PBSCT the viral load increased again. Without further VSTs, however, it started to decrease on day 269. From day 566 onwards, CMV viral load remained undetectable.
Cellular CMV immunity was determined by IFN-γ ELISpot, based on the stimulation of 200,000 peripheral blood mononuclear cells (PBMCs) with pp65 and IE-1 CMV proteins (T-Track® CMV-Assay, Lophius Biosciences GmbH, Regensburg, Germany), as described recently . This CE-approved assay measures CMV-specific responses of both CD4+ and CD8+ T lymphocytes within 19 h. Spot numbers were determined by an ELISpot plate reader (AID Fluorospot, Autoimmun Diagnostika GmbH, Strassberg, Germany). The median spot number of duplicate cultures was considered for further analysis and values of negative controls were subtracted from CMV-specific values (spots increment). The sister's PBMC showed strong CMV immunity (423 CMV pp65 and 16 CMV IE-1 spots increment), whereas CMV immunity was very weak in the patient prior to VSTs (12 CMV pp65 and 5 CMV IE-1 spots increment; Fig. 1b). After VSTs, CMV pp65-specific spots increased until day 713 (367 spots increment). However, a minimum on day 110 and day 144 could be observed. CMV IE-1-specific spots reached a maximum on day 123 (19.5 spots increment) and then started to decrease. Taken together, viral control by VSTs could be assumed for 41 days after VSTs.
Moreover, CMV-specific T cells were determined by multimer analysis as previously described [6, 9], using HLA-A*02- and HLA-B*07-restricted peptides (Fig. 1c). After VSTs, the percentage of CMV pp65- and IE-1-specific IFN-γ+ CD8+ T cells increased up to 0.84 and 0.53%, respectively.
Increased viral replication was always followed by low numbers of leukocytes and neutrophils (Fig. 1d), which may be due to myelotoxic antiviral drugs (valganciclovir and foscarnet).
Finally, we analyzed the origin of the CMV-specific cells that provided long-term control of the virus. On day 713 after transplantation, patient PBMCs were stimulated with the CMV pp65 protein, using the same conditions as for the ELISpot. Patient PBMC without CMV stimulation served as control. Hematopoietic chimerism testing was performed using the KMRtype® and KMRtrack® Chimerism Monitoring Reagents (GenDx, Utrecht, the Netherlands) according to the manufacturer’s recommendations, determining bi-allelic insertion–deletion polymorphism . The input DNA for each reaction was 100 ng, resulting in a detection limit of 0.06%. We found 100% donor chimerism in the CMV-specific cells as well as in the cells without CMV stimulation. Thus, all cells displayed the genetic markers of the unrelated donor. No genetic markers of the sister who donated the VSTs or of patient origin were observed. Moreover, XY-FISH analysis  was performed nearly every month after PBSCT and showed 100% donor chimerism in the peripheral blood of the patient, e.g., immediately prior to VSTs, on day 48 and day 917 after VSTs. On day 54 after VSTs also CD3+ T cells (without CMV stimulation) were tested by XY-FISH analysis, yielding 100% donor cells. A disease-specific marker, a mutation of nucleophosmin 1 (NPM1) , which was present prior to transplantation, remained undetectable throughout the whole follow-up after PBSCT. Most likely, CMV disease was controlled early by the VSTs and later by CMV-specific immunity that was established from the unrelated donor-derived immune system.
Overall, side effects of VSTs were moderate. On day 144 after PBSCT (day 48 after VSTs) the patient suffered from acute GvHD of the skin (stage 2) and of enteral GvHD (stage 2), i.e., grade 2. On day 199 after PBSCT she presented with moderate, histologically confirmed chronic GvHD with sclerodermic areas, exudative enteropathy, and malabsorption with hypoproteinemia edema, which improved after increasing the dose of prednisone. Starting on day 214 after PBSCT the dose of mycophenolate mofetil and prednisone could be tapered. Since day 287 the treatment with prednisone could be stopped and since day 370 the patient is without any immunosuppression. Moreover, at day 356 after PBSCT, lymphocyte subpopulations reached the level of healthy controls. Currently, 3 years after VSTs, the patient is in good health and is able to work. Thus, alloreactivity that could have been induced by VSTs was treated successfully.
In summary, our data indicate that CMV replication could be intermittently controlled by VSTs from a CMV-positive donor. Thus, if antiviral therapy fails VSTs may represent a suitable treatment alternative. Presumably, a second infusion of VSTs ~1 month after the first could have prevented the subsequent re-appearance of the virus. The course of the viral load (Fig. 1a) indicates that it took more than 6 days until the VSTs led to a decrease of viral replication. Since the patient and her sister who donated the VSTs were HLA-haploidentical, alloresponses toward the foreign HLA haplotype may have limited the lifespan of the VSTs. However, in a different solid organ transplant recipient who received EBV-VSTs from a 5/10 HLA-matched donor, these donor T cells could be detected for 1 year, i.e., the end of the current follow-up. According to our previous data , the transfer of antiviral immunity may last longer if donor and recipient were HLA-identical. In HLA-identical donor/recipient pairs donor-derived hepatitis B virus-specific immunity was detectable for up to 5 years after hematopoietic stem cell transplantation.
In conclusion, in selected cases the infusion of VSTs should be considered to control CMV replication. The optimal frequency of VSTs' application may depend on the histocompatibility of donor and recipient.
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We are grateful to Monika Huben, Anja König, Nicole Neuman, Martina Praast, and Dörthe Rokitta for their excellent technical assistance.
D.W.B., B.E.V., M.L., R.B., and M.K. were involved in the conception and design of the study. M.K., N.K.S., V.K., and D.W.B. provided the samples. M.L., M.K., B.M.K., M.F., and A.H. were involved in the collection and assembly of data. B.E.V., B.M.K., and R.B.L. were involved in the donor selection, quality control of the CMV-VSTs, and the final release of the T-cell product. M.L. and M.K. were involved in data analysis and interpretation. M.L., M.K., and P.H.O. wrote the manuscript. All the authors gave final approval of the manuscript.
Conflict of interest
The authors declare that they have no conflict of interest.
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Lindemann, M., Eiz-Vesper, B., Steckel, N.K. et al. Adoptive transfer of cellular immunity against cytomegalovirus by virus-specific lymphocytes from a third-party family donor. Bone Marrow Transplant 53, 1351–1355 (2018). https://doi.org/10.1038/s41409-018-0209-2
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