Automated Cell Enrichment of Cytomegalovirus-specific T cells for Clinical Applications using the Cytokine-capture System

1Division of Pediatrics, U.T. MD Anderson Cancer Center, 2Stem Cell Transplantation and Cellular Therapy, U.T. MD Anderson Cancer Center
* These authors contributed equally
Published 10/05/2015
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Immunology and Infection

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Summary

The goal of this protocol is to manufacture pathogen-specific clinical-grade T cells using a bench-top, automated, second generation cell enrichment device that incorporates a closed cytokine capture system and does not require dedicated staff or use of a GMP facility. The cytomegalovirus pp65-specific-T cells generated can be directly administered to patients.

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Kumaresan, P., Figliola, M., Moyes, J. S., Huls, M. H., Tewari, P., Shpall, E. J., et al. Automated Cell Enrichment of Cytomegalovirus-specific T cells for Clinical Applications using the Cytokine-capture System. J. Vis. Exp. (104), e52808, doi:10.3791/52808 (2015).

Abstract

The adoptive transfer of pathogen-specific T cells can be used to prevent and treat opportunistic infections such as cytomegalovirus (CMV) infection occurring after allogeneic hematopoietic stem-cell transplantation. Viral-specific T cells from allogeneic donors, including third party donors, can be propagated ex vivo in compliance with current good manufacturing practice (cGMP), employing repeated rounds of antigen-driven stimulation to selectively propagate desired T cells. The identification and isolation of antigen-specific T cells can also be undertaken based upon the cytokine capture system of T cells that have been activated to secrete gamma-interferon (IFN-γ). However, widespread human application of the cytokine capture system (CCS) to help restore immunity has been limited as the production process is time-consuming and requires a skilled operator. The development of a second-generation cell enrichment device such as CliniMACS Prodigy now enables investigators to generate viral-specific T cells using an automated, less labor-intensive system. This device separates magnetically labeled cells from unlabeled cells using magnetic activated cell sorting technology to generate clinical-grade products, is engineered as a closed system and can be accessed and operated on the benchtop. We demonstrate the operation of this new automated cell enrichment device to manufacture CMV pp65-specific T cells obtained from a steady-state apheresis product obtained from a CMV seropositive donor. These isolated T cells can then be directly infused into a patient under institutional and federal regulatory supervision. All the bio-processing steps including removal of red blood cells, stimulation of T cells, separation of antigen-specific T cells, purification, and washing are fully automated. Devices such as this raise the possibility that T cells for human application can be manufactured outside of dedicated good manufacturing practice (GMP) facilities and instead be produced in blood banking facilities where staff can supervise automated protocols to produce multiple products.

Introduction

Hematopoietic stem-cell transplantation (HSCT) 1 can be combined with adoptive T-cell therapy to improve graft-versus-tumor effect and to provide immunity to opportunistic infections2. Generation of antigen-specific donor-derived T cells for infusion has historically required skilled personnel and use of specialized facilities that are GMP-compliant. The delivery of such T cells has resulted in resolution of opportunistic infections3 as well as treating the underlying malignancy4. Recently, investigators have demonstrated that the adoptive transfer of only few thousand virus-specific T cells (~ 1 x 104 –  2.5 x 105 cells/kg recipient body weight) can successfully treat opportunistic CMV infections after allogeneic HSCT5-9. A limited number of GMP facilities with associated skilled manufacturing requirements and the high cost associated with cell production has, however, restricted patient access to promising T-cell therapies10. One approach to isolating antigen-specific T cells is based on the CCS using a bi-specific reagent to recognize CD45 and IFN-γ. As is shown, this methodology can be used to generate clinical-grade CMV-specific T cells employing an automated cell enrichment CCS device (Figure 1B).

CMV-specific T cells are generated by incubating overlapping peptides from CMV pp65 antigen with leukapheresis total nuclear cells (TNC) from CMV-seropositive donors. These peptides, displayed in the context of human leukocyte antigen (HLA), activate the CMV pp65-specific T cells within the TNC to secrete IFN-γ. These T cells can then be “captured” and magnetically separated. The operation of the first-generation cell enrichment device (Figure 1A) required personnel skilled in cell culture under GMP conditions, and coordination of staff to undertake the multiple steps necessary to generate a “captured” product.

The procedure typically required 10 to 12 hr of continuous operation, and therefore personnel likely need to work over two shifts in the GMP facility. These constraints are now obviated by the implementation of a second-generation device (shown in Figure 1B). This device undertakes magnetic enrichment, similar to the first generation device, but automates other aspects of the CCS in an unbreached approach. This significantly reduces the burden on the GMP team as most of the steps can be accomplished unattended by staff. Furthermore, since the device operates as a closed system, the antigen-specific T cells can be captured and processed on the benchtop except the steps involved in leukapheresis isolation and preparation of materials before starting the instrument. Details of the complete instrumentation and functionality of this second-generation cell enrichment device have been published11.

Here, we describe the steps to enrich CMV pp65-specific T cells from a steady-state apheresis product using the automated cell enrichment CCS system. Once isolated, these CMV-specific T cells may be immediately infused into a patient.

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Protocol

1. Preparation of Materials under Sterile Conditions (See Materials and Equipment Table)

  1. Prepare 3 L of PBS/EDTA buffer supplemented with human serum albumin (HSA) to a final concentration of 0.5% (w/v).
  2. Prepare 1 L bag of clinical grade 0.9 % sodium chloride (NaCl) solution and 2 L of GMP grade cell culture medium.
  3. Prepare 60 nmol of CMV-specific peptide antigen cocktail by reconstituting one vial of CMV pp65 with 8 ml of sterile water.
  4. Transfer CMV pp65 peptide cocktail into a 50 ml volume freezing bag using a Luer/Spike interconnector and clamp with locking forceps to avoid subsequent distribution of cocktail into the tubing set. Open cell enrichment tubing set (TS 500) under sterile conditions.
  5. Using sterile tubing welder, connect peptide cocktail freezing bag into tube connection for valve 2 of tubing set TS 500. Do not open the peptide cocktail bag clamp at this time.
  6. Remove 1 x 109 TNC from starting cellular product and suspend in PBS/EDTA buffer containing 2.5% HSA to a total volume of 50 ml. Inject the cellular product into a 150 ml transfer bag.

2. Preparation and Use of Automated Cell Enrichment System (See Materials and Equipment Table)

  1. Switch on the cell enrichment system (Figure 1B) and select the program “ CCS_IFN-γ Enrichment”. Observe a user interface showing screens with instructions and pictures guiding the operator through the procedure.
  2. Enter the parameter “Operator” and “Tubing Set P/N No”. Next, install the Tubing Set 500 to automated cell enrichment device as per instructions displayed on the interactive monitor screen.
  3. Follow the step by step instructions displayed on the screen to connect the medium and buffers to the device. Record catalogue number and lot number of the reagents before connecting to the instrument.
  4. After the final check of the tubing set, open the clamp of the peptide cocktail bag. Open the medium bag and initiate automatic priming of the tubing set.
  5. After the priming step is completed, supplement HSA (2.5%) into NaCl buffer in the reservoir bag (200 ml) with the help of the sterile tubing welder. Transfer the starting cellular product into the “Application bag” using the sterile tubing welder.
  6. Connect CCS (IFNγ) reagents into respective tubing via adapters. Enter preferred time to collect a fraction of cellular material before enrichment process. Review and verify the accuracy of all data/parameters entered. Start the process.
  7. Before the start of automated cell enrichment process, remove the Quality Control Bag (QCB, original fraction (ori) contains approximately 1.3 ml out of 100 ml chamber content diluted with PBS/EDTA buffer). Seal the QCB, weigh, and store at 4 °C.
  8. Start the enrichment process. At the end of the process, target cells will be eluted with an approximate volume of elution buffer from the reservoir bag.
  9. Seal the Non Target Cell Bag (NTCB, negative fraction = neg) and Target Cell Bag (TCB, positive fraction = pos) and weigh each bag. The weights will be used later for calculation of the cell numbers.
  10. Immediately after the enrichment procedure collect two aliquots per fraction for flow cytometry analysis, and store the rest of the samples at 4 °C. Use one sample aliquot for cell count determination and the other sample aliquot for the enrichment performance analysis (Table 1).
  11. Remove the tubing set from the cell enrichment instrument. Transfer the log file to a USB drive for future use.
    NOTE:  All reagents should be prepared under sterile conditions. The use of a Biosafety type II hood is highly recommended. Use steady-state apheresis cellular product (non-mobilized) isolated from a healthy CMV-seropositive donor to enrich CMV antigen specific T cells. Only FDA licensed HSA should be used. The buffer for cell preparation should be kept at  +19 °C to +25 °C as lower or higher ambient temperatures will result in reduced purity and a reduced yield of the target cells.

3. Cell Count Determination

  1. Take the aliquots of QCB , NTCB and TCB for cell counts as shown in Table 1. Add CD45-VoBlue to each aliquot (titer 1:11) and incubate in the dark for 10 min at 4 °C.
  2. Add 1.5 ml freshly prepared red blood cell lysis solution (1x) to the original fraction and negative fraction, 450 µl freshly prepared red blood cell lysis solution to the positive fraction, and incubate all fractions for 15 min at RT.
  3. Just prior to analysis, add propidium iodide to a final concentration of 1 µg/ml (1:100 dilution of 100 µg/ml). Use automatic cell counter to determine cell count and viability. Use cell counter device recommended software for flow cytometry analysis. Determine the absolute counts of leukocytes for original, negative and positive fractions.
     NOTE: The cell count of viable leukocytes per ml of the samples taken for cell count analysis is determined using the cell analyzer recommended software.
  4. Set the region as shown in Figure 2 (region 5, viable leukocytes). Use the following gating strategy to determine cell count. A viable leukocyte in original fraction is shown in Figure 2.
  5. The indicated regions (Figure 2, 1-6) are hierarchically as follows:
    1: Time gate → 2: Single cells → 3: CD45+ cells → 4: Leukocytes (debris excluded) → 5: Viable leukocytes → 6: Viable lymphocytes
  6. Repeat the same steps to determine cell counts for negative and positive fractions. Calculate the cell count of the whole fraction by considering the diluent factor of the sample and total volume of the fraction (Table 2).

4. Examination of the Separation Performance

  1. Wash the aliquots of QCB, NTCB and TCB fraction cells with pre-chilled PBS/EDTA buffer/0.5% AB serum. Centrifuge the cells at 300 x g for 5 min at 4 °C and aspirate the supernatant.
  2. Resuspend cells in 100 µl antibody-fluorochrome staining mixture containing: CD3-FITC, CD4-APC, CD8-APC-Vio770, CD14-PerCP, CD20-PerCP, CD45-VioBlue and anti-IFNγ-PE (titer 1:11) and incubate in the dark for 10 min at 4 °C.
  3. Add 1 ml freshly prepared red blood cell lysis solution (1x) and incubate for 15 min at RT Centrifuge at 300 x g for 5 min at 4 °C and aspirate the supernatant Resuspend the cells in an adequate volume of PBS/EDTA Buffer/ 0.5% AB Serum.
  4. Add propidium iodide to a final concentration of 1 µg/ml just prior to analysis (1:100 dilution of 100 µg/ml). Perform flow cytometry analysis to evaluate the purity of the sample.
  5. Use the following gating strategy to calculate the CD3+ T cells in viable leukocyte Gating strategy for determining the CD3+ T cells is shown in positive fraction after CCS enrichment process. The indicated regions (Figure 3A and 3B, 1-6) are hierarchically linked as follows:
    1: Time gate → 2: Single cells → 3: CD45+ cells → 4: Cells (debris excluded) → 5: Viable leukocytes → 6: Viable CD3+ cells population
  6. Determine the frequencies of CD4+, CD8+, CD4+ IFN-γ+ and CD8+ IFN-γ+ T cells after CCS enrichment process (Table 2).
  7. Use the gating strategy to determine the frequencies of CD4+, CD8+, CD4+ IFN-γ+ and CD8+ IFN-γ+ T cells shown below for an original and enriched (captured) positive fraction after CCS process. The indicated regions are hierarchically linked and named as follows:
    1: Time gate → 2: Single cells → 3: CD45+ cells → 4: Cells (debris excluded) → 5: Viable Leukocytes → 6: Viable CD3+ cells → 7: CD4+ cells → 7a: CD4+ IFN-γ+ cells (box) → 8: CD8+ cells → 8a: CD8+ IFN-γ+ cells (box)  

NOTE: The first 6 indicated regions of the hierarchy links are the same as Figure 3, (1-6) and last 2 regions are shown in Figure 4 (6-8a).

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Representative Results

In this study, an automated cell enrichment CCS System was used for automated production of CMV pp65-specific T cells. CMV-specific T cells were enriched from three apheresis cell products. The steady-state apheresis product was harvested over 2 hr from a CMV-seropositive donor and generated 1010 total nuclear cells (TNC). 109 TNC were then activated with CMV pp65-derived peptides (60 nmol) for 4 hr and the IFN-γ secreting T cells were isolated using the CCS on the automated cell enrichment device. An operator was needed at the beginning of the experiment to load all the reagents and tubing sets. Setup of the system from the initial unpacking to starting the device took approximately 60 to 120 min. Note, the machine can then be programmed to start after the reagents and tubing sets are loaded thereby enabling the machine to operate unattended (such as overnight). The operator was needed again after 15 hr to perform characterization of the final products for cell purity and viability. After enrichment the cell supernatant was screened for mycoplasma and endotoxin presence. After validation, the processed cells can be infused directly into patients or cryopreserved for later applications.

Cell counts were determined following cell counting standard practices using the formulas given in Table 2. Each type of cell count was repeated three times and the results were expressed as mean total cell counts with standard deviation (SD). Reports were then analyzed using cell analyzer recommended software. Gating to determine viable leukocytes and lymphocytes is shown in Figure 2. The data for the cell counts are presented in Table 3. The gating strategy used to determine IFN-γ+ T cells is shown in Figure 3 and Figure 4. Before enrichment, the viability of cells was routinely > 95%. After enrichment, the viability of cells was < 50%. The absolute count of IFN-γ+ T cells was assessed before and after the enrichment process. The total number of IFN-γ+ T cells before enrichment was 1.14 x 106 ± 0.35 x 106 as derived from 109 starting TNC, and after enrichment was 3.09 x 105 ± 1.70 x 105 IFN-γ+ T cells. There were 0.16 ± 0.18% IFN-γ+ CD4+ T cells present prior to processing and this increased to 47.5 ± 34.7% after enrichment. The purity percentage of CD8+ IFN-γ+ T cells prior to capture was 0.47 ± 0.1%, and increased to 90.3 ± 1.7% after enrichment (Table 3 and 4). Sample recovery in the captured (positive) fraction was 32.9 ± 15.7% for CD4+ T cells and 31.8 ± 13.2% for CD8+ T cells based on measurement of IFN-γ+ T cells in the starting population (Table 4). These data indicate that both CD4+ and CD8+ CMV pp65-specific T cells can be harvested automatically in a manner suitable for their human application.

Figure 1
Figure 1. Enrichment of CMV-specific T cells using CCS system. (A) Multiple processing steps involved in the first generation cell enrichment device are handled by skilled professionals. (B) Most of the processing steps, except initial tubing setup, are automated in the second generation cell enrichment device which saves 10 - 12 hr of operator handling time in comparison with the first generation device. The enriched CMV-virus-specific T cells are characterized by the flow cytometry cell analyzer. Please click here to view a larger version of this figure.

Figure 2
Figure 2. Gating strategy used to determine viable T cells. The numbers 1 through 6 indicate the corresponding gating hierarchy domain in the figure. (1) Setting up time gate, (2) Removing doublet cells by plotting FSC-height against FSC-area (3) Identifying CD45+ cells, (4) Removing cell debris (5) Selecting viable leukocytes and (6) Viable lymphocytes from original population by propidium iodide staining. Please click here to view a larger version of this figure.

Figure 3
Figure 3. Gating strategy used to determine CD3+ T cells. Flow cytometry blot analysis of CD3+T cells before (3A) and after (3B) cell enrichment is shown here. It is crucial to determine how many CMV-specific peptide activated T cells are present in the samples before and after the enrichment process. In this figure, numbers 1 through 6 indicate the corresponding cell population either by size or stained by a specific antibody. (1) Setting up time-gate, (2) Removing doublet cells, (3) Selecting CD45+ cells, (4) Removing cell debris (5) Selecting viable leukocytes and (6) Viable CD3+ lymphocytes. Propidium Iodide staining was performed to remove dead cells. Please click here to view a larger version of this figure.

Figure 4
Figure 4. Gating strategy used to determine purity of IFN-γ+ T cells. Activated CMV-specific T cells are critical for controlling CMV infection, so IFN-γ+ expression on T cells were used for gating. IFN-γ+ T cells among CD4+ and CD8+ subsets (A) before enrichment and (B) after enrichment. (7/8) Percentage of CD4+ T cells and CD8+ T cells is shown in A and B. (7a) Percentage of CD4+IFN-γ+ T cells is shown in the square box (a) within the gating area and similarly for CD8+IFN-γ+ T cells in (8a). “T” represents % of cells in total population and “#” represents % of cells in gated population. Propidium iodide staining was performed to gate-out dead cells. Please click here to view a larger version of this figure.

Table 1
Table 1. Volumes of the fractions used for cell staining strategy.

Table 2
Table 2. Formulas used for calculation of CD4+ IFN-γ+ T cells after enrichment process. Calculation of CD8+ IFN-γ+ T cells is performed similarly.

Table 3
Please click here to view a larger version of this table.
Table 3. Total cell counts in T-cell subsets before and after enrichment. Sample #1 results are shown here.

Table 4
Table 4. Purity and recovery of IFN-γ+ T cell before and after enrichment process of sample #1.

Table 5
Table 5. Cell sorting strategies used in the isolation of clinical grade CMV-antigen specific T cells.5

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Discussion

Adoptive T-cell therapy has emerged as a viable option to treat B-cell malignancies4. Its therapeutic potential is dependent on infusing the desired number of target antigen specific T cells that lack replicative senescence2. This can be achieved by sorting out a pure population of antigen specific T cells from expanded T cells in compliance with current good manufacturing practices. Two sorting procedures are widely used, namely, fluorescence-activated cell sorting (FACS) and magnetic activated cell sorting (MACS) to generate CMV antigen specific T cells as we have reviewed recently5.   The advantage of using one strategy over the other is outlined in Table 5. MACS technology offers the highest cell enrichment purity in a cheaper and faster way compared to FACS technology. In addition usage of disposable columns and reagents for cell enrichment totally prevents sample to sample contamination making it easier to apply in clinical settings. The semi-automated cell enrichment device is labor intensive and time consuming so development of the automated cell enrichment device was necessary to feed clinical demand.

The automated cell enrichment CCS device is a versatile instrument for isolating clinical grade cells such as hematopoietic stem cells, somatic stem cells, as well as T cells for adoptive transfer. This automated device integrates cell processing, including fractionation of starting material, cell washing, cell separation, cell culture, and final product formation in a single-use GMP-compliant disposable unit. The closed system reduces clean-room requirements and minimizes operator involvement for maintaining GMP facilities. Automation reduces the time required for an operator to be present thereby decreasing the cost associated with these laboratory procedures.

To validate the automated cell enrichment device, as compared with the semi-automated cell enrichment device, we isolated CMV-specific T cells after incubating CMV pp65-derived peptides with an apheresis product. GMP grade CMV pp65-derived peptide cocktail (e.g. PepTivator) is a peptide pool that consists mainly of 15 mer-peptides with 11 amino acids overlap, covering the complete sequence of the pp65 protein of human cytomegalovirus. Sample recovery yield was ~ 0.3 x 106 CD3+IFN-γ+ T cells from 109 TNC. Clinical studies have demonstrated that infusing a few thousand CMV-specific T cells as prophylactic treatment for patients (~ 360 to 4,000 cells/kg body weight) undergoing HSCT resulted in protection against CMV. For example, in order to treat CMV in clinical trials using this technology, a 70 kg adult and a 30 kg child apparently require only 0.26 – 3.0 x 105 and 1 x 104 – 1.2 x 105 cells, respectively, of viral-specific T cells12-15. In general, the number of live/dead cells is comparable to the semi-automated cell enrichment device, as shown by Feuchtinger et al.13. A higher number of viable cells would also be acceptable as this would be associated with enough material for different infusions. However, more viable cells in general also means more cells other than T cells (B, NK, etc). Our data demonstrate that CMV-specific T cells generated on the automated cell enrichment system resulted in clinically-appealing numbers of CMV-specific T cells that may then be infused after allogeneic HSCT.

CMV infection can be a major problem after HSCT resulting in both increased morbidity and mortality. Furthermore, CMV infection is associated with increased costs, despite recent progress in early diagnosis and early treatment with anti-viral drugs16. The current treatment using ganciclovir and foscarnet can lead to toxicity in medically-fragile recipients of HSCT. The add-back of donor-derived CMV-specific T cells has been demonstrated to prevent and treat opportunistic infections in recipients of allogeneic HSCT9. This approach to adoptive immunotherapy has also been applied to help restore immunity to other pathogens, such as Epstein-Barr virus (EBV), adenovirus8, and Aspergillus3 by incubating mononuclear cells (MNC) with respective antigen derived clinical grade peptide cocktail reagents (Materials and equipment’s table). Recently, investigators have safely infused third-party pathogen-specific T cells that are matched with at least one HLA allele in the recipient presenting immunodominant peptide17. The automated cell enrichment CCS system may also be used to generate third-party T cells for off-the-shelf applications that will be useful when the donor is CMV-seronegative or unavailable, such as the case with donors for allogeneic umbilical cord blood transplantation.

In summary, we demonstrate the utility of an automated cell enrichment device to generate CMV-specific T cells based on automation of CCS. We believe this device has the potential to lower the threshold for clinical teams to infuse pathogen-specific, as well as tumor-specific, T cells in immunocompromised patients.

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Disclosures

Both MD Anderson Cancer Center and Dr. Cooper have a financial interest in ZIOPHARM Oncology, Inc., and Intrexon Corporation. On May 7, 2015, Dr. Cooper was appointed as the Chief Executive Officer at ZIOPHARM Oncology.  Dr. Cooper is now a Visiting Scientist at MD Anderson. Dr. Cooper founded and owns InCellerate, Inc. He has patents with Sangamo BioSciences with artificial nucleases. He consults with Targazyme, Inc. (formerly American Stem cells, Inc.), GE Healthcare, Ferring Pharmaceuticals, Fate Therapeutics, Janssen Pharmaceuticals, and Bristol-Myers Squibb. He is on the Scientific Advisory Board of Cellectis. He receives honoraria from Miltenyi Biotec.

Acknowledgements

We thank Miltenyi Biotec, Germany for providing reagents and CliniMACS Prodigy equipment for evaluation studies. We thank George T. McNamara (Pediatric department, MD Anderson Cancer Center) for proof reading the manuscript. Grant support: Cancer Center Core Grant (CA16672); RO1 (CA124782, CA120956, CA141303; CA141303); R33 (CA116127); P01 (CA148600); Burroughs Wellcome Fund; Cancer Prevention and Research Institute of Texas; CLL Global Research Foundation; Estate of Noelan L. Bibler; Gillson Longenbaugh Foundation; Harry T. Mangurian, Jr., Fund for Leukemia Immunotherapy; Institute of Personalized Cancer Therapy; Leukemia and Lymphoma Society; Lymphoma Research Foundation; MDACC’s Sister Institution Network Fund; Miller Foundation; Mr. Herb Simons; Mr. and Mrs. Joe H. Scales; Mr. Thomas Scott; National Foundation for Cancer Research; Pediatric Cancer Research Foundation; William Lawrence and Blanche Hughes Children's Foundation.

Materials

Name Company Catalog Number Comments
CliniMACS PBS/EDTA Buffer 3 L bag Miltenyi Biotec GmbH 700-29
CliniMACS Prodigy Tubing Set TS 500 Miltenyi Biotec GmbH 130-097-182
5 L waste bag Miltenyi Biotec GmbH 110-004-067
CliniMACS Cytokine Capture System (IFN-gamma) Miltenyi Biotec GmbH 279-01
Albumin (Human) 25%  Grifols 58516-5216-2
Luer/Spike Interconnector Miltenyi Biotec GmbH 130-018-701
0.9 % NaCl Solution (1 L) Miltenyi Biotec GmbH
MACS GMP PepTivator HCMV pp65 Miltenyi Biotec GmbH 170-076-109
Water for injections Hospira, inc, Lake Forest, IL NDC-0409-4887-10
MILLEX GV Filter Unit 0.22 μm  Millipore SLGV033RB
TexMACS GMP Medium 2 L bag Miltenyi Biotec GmbH 170-076-306
Transfer Bag, 150 ml (for cellular starting material) Miltenyi Biotec GmbH 130-018-301
CryoMACS Freezing Bag 50 Miltenyi Biotec GmbH 200-074-400
60 ml Syringes, sterile BD, Laagstraat, Temse, Belgium 309653
CMV sero positive apheresis product Key Biologics, LLC, Memphis
Flow Cytometry Materials Manufacturer Catalog number
AB Serum, GemCell Gemini Bio-Products, West Sacramento, USA 100-512
CD3-FITC Miltenyi Biotec GmbH 130-080-401
CD4-APC Miltenyi Biotec GmbH 130-098-033
CD8-APC-Vio770 Miltenyi Biotec GmbH 130-098-065
CD14-PerCP Miltenyi Biotec GmbH 130-098-072
CD20-PerCP Miltenyi Biotec GmbH 130-098-077
CD45-VioBlue Miltenyi Biotec GmbH 130-098-136
aIFN-γ-PE, human Miltenyi Biotec GmbH 130-097-940
CD3-PE Miltenyi Biotec GmbH 130-091-374
Propidium Iodide Solution (100 µg/ml) Miltenyi Biotec GmbH 130-093-233
Equipment Manufacturer Catalog Number
CliniMACS Prodigy Device  Miltenyi Biotec GmbH 200-075-301
Software V1.0.0.RC
MACSQuant Analyzer 10 Miltenyi Biotec GmbH 130-096-343
Software 2.4
Centrifuge 5415R  Eppendorf AG 22331
Cellometer K2 Nexelom Bioscience, Lawrence, MA LB-001-0016
Sterile tubing welder SCDIIB Terumo Medical Corp., Elkton, MA 7811

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References

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  9. Meij, P., et al. Effective treatment of refractory CMV reactivation after allogeneic stem cell transplantation with in vitro-generated CMV pp65-specific CD8+ T-cell lines. J Immunother. 35, (8), 621-628 (2012).
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  12. Brestrich, G., et al. Adoptive T-Cell Therapy of a Lung Transplanted Patient with Severe CMV Disease and Resistance to Antiviral Therapy. Am J Transplant. 9, (7), 1679-1684 (2009).
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  14. Peggs, K. S., et al. Directly selected cytomegalovirus-reactive donor T cells confer rapid and safe systemic reconstitution of virus-specific immunity following stem cell transplantation. Clin Infect Dis. 52, (1), 49-57 (2011).
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  17. Leen, A. M., et al. Multicenter study of banked third-party virus-specific T cells to treat severe viral infections after hematopoietic stem cell transplantation. Blood. 121, (26), 5113-5123 (2013).

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