Reconstitution of FOXP3⁺ regulatory T cells (T_regs) after CD25-depleted allotransplantion in elderly patients and association with acute graft-versus-host disease

Severe graft-versus-host disease (GvHD) in the early posttransplantation period limits the success of allogeneic stem-cell transplantation (SCT) and requires powerful immunosuppression that deprives the allotransplant of its full graft-versus-leukemia (GvL) potential.¹  Selective depletion (SD) of host-reactive donor T cells from lymphocyte products is a promising strategy to prevent severe, acute GvHD (aGvHD) while maintaining useful donor-derived immune functions such as GvL effects and antiviral immunity.²  A variety of experimental approaches for the ex vivo removal of alloreacting lymphocytes has been proposed targeting surface expression of activation-associated molecules such as CD25,^3-12  CD69,^9,10,12,13  CD71,¹²  HLA-DR,¹²  and CD137,¹⁴  alterations in the multidrug resistance pump p-glycoprotein,^15,16  or proliferation.^17,18  Targeted T cells can be removed or eliminated by using immunomagnets,^8,9,12,14  immunotoxins,^3-7,11,19  flow sorting,^17,18  induction of apoptosis,^19,20  or photodepletion techniques.^15,16  Most clinical experience concerns SD techniques using an anti-CD25 immunotoxin (CD25-IT) to target the α-chain of the interleukin-2 (IL-2) receptor (CD25) to eliminate ex vivo–activated donor lymphocytes.^21-23  We recently reported a 46% grade II to IV and 12% grade III to IV aGvHD incidence in 16 elderly patients with advanced hematologic malignancies who were treated with selectively CD25-depleted allografts from HLA-matched siblings.²³  Because CD25 is not exclusively expressed on effector T cells (T_effs), but also on a subset of CD4⁺ regulatory T cells (T_regs) that suppress alloresponses and protect against GvHD,^24-32  there is a concern that the concurrent removal of T_regs could have increased the risk of GvHD in our series.

While the phenotypic distinction between T_regs and T_effs has been challenging in the past, especially in humans when based on CD25 alone, nowadays availability of monoclonal antibodies directed against the forkhead box protein 3 (FOXP3) make it possible to identify a subset of CD25⁺ FOXP3⁺ T_regs from CD25⁺ FOXP3⁻ T_effs.^33-36  FOXP3 encodes a forkhead/winged helix transcription factor and was identified as a key regulator required for the development and functional activity of T_regs.^33-35  To characterize T_reg recovery after SD transplantation, we therefore retrospectively measured T_regs in transplant products and in 16 of our patients receiving SD transplants in the first 3 months after SCT.²³  We found that T_reg recovery was prompt and may in part be derived from a CD25⁻ T_reg content persisting in the SD product.

Patients and their HLA-identical sibling donors were treated on the National Institutes of Health protocol 01-H-0162, approved by the National Heart, Lung, and Blood Institute Review Board. All patients and donors provided written informed consent in accordance with the Declaration of Helsinki before enrollment. Based on sample availability, T_regs were analyzed in peripheral blood samples from 16 patients (15 previously reported²³ ) with advanced hematologic malignancies (before transplantation; 30, 60, and 90 days after transplantation), 13 of their respective stem cell donors, and 10 of the selectively CD25-depleted (SD) products. Table 1 outlines patient demographics, conditioning regimens, stem and T-cell doses, transplantation outcomes, and sample availability. As analyses for FOXP3 gene expression by quantitative reverse-transcription–polymerase chain reaction (qRT-PCR) preceded those of FOXP3 intracellular staining in time, not all samples were available for both assays. All samples analyzed came from patients prior to donor lymphocyte infusions (DLIs) or GvHD treatment with anti-CD25.

The transplantation approach and the selective depletion procedure have been described previously.^5,23 Figure 1 provides a schematic overview. Briefly, older patients with advanced hematologic malignancies and a suitable HLA-matched sibling stem cell donor were eligible for the study. All patients received a preparative regimen consisting of fludarabine (125-180 mg/m²) and one of the following alkylating agents: cyclophosphamide (120 mg/kg), melphalan (140 mg/m²), or infusional busulfan (6.4 mg/kg). Donors received granulocyte colony-stimulating factor (G-CSF) 10 μg/kg per day subcutaneously. Mobilized peripheral blood stem cells (PBSCs) were collected by leukapheresis on day 5, and again on days 6 and 7 if necessary, to obtain a target dose of at least 3 × 10⁶ CD34⁺ cells/kg. The Isolex 300i immunomagnetic system (Nexell Therapeutics, Irvine, CA) was used to perform a combined positive CD34⁺ cell selection and negative selection of T cells according to the manufacturer's instructions, which results in a PBSC product with approximately 5 logs of T-cell depletion.

The unadsorbed, T-cell–rich fraction remaining after CD34 selection was the source of donor lymphocytes for the SD procedure. Recipient stimulator cells were generated from immunomagnetically selected and expanded T lymphocytes. Stimulator cells were thawed, gamma irradiated (2500 cGy), and added to responder cells at a ratio of 1 to 1 to a final concentration of 5 × 10⁶ cells/mL. Responder and irradiated stimulator cells were cocultured for 72 hours at 37°C in 7% CO₂. Alloreactive responder cells expressing the activation marker, CD25, were targeted by the addition of the anti-CD25 immunotoxin, RFT5-SMPT-dgA, which consists of the anti-CD25 murine MAb, RFT5 (IgG1), coupled to the deglycosylated ricin A chain (dgA) via the heterobifunctional cross-linker SMPT (Pierce, Rockford, IL). RFT5-SMPT-dgA was added to the culture at 24 (2 μg/mL) and 48 (1 μg/mL) hours, along with the enhancing agent, ammonium chloride (0 mM; Sigma, St Louis, MO). At the completion of the 72-hour coculture, cells were harvested and cryopreserved. On the day of transplantation (day 0), both products (stem cells and SD T cells) were thawed and infused consecutively. Patients received cyclosporine alone for GvHD prophylaxis starting on day − 5. In the absence of aGvHD, immunosuppression was tapered and then withdrawn by day + 100.

Cells were phenotypically analyzed by 5-color flow cytometry using CD3-PE-Cy7 (clone SK7), CD4-PerCP (clone SK3), CD25-PE (clone M-A251), and CD27-FITC (clone L128) antibodies for surface staining (all from BD Biosciences, San Diego, CA). Intracellular staining of FOXP3 (eBioscience, San Diego, CA) was performed after surface staining, fixation, and permeabilization according to the manufacturer's recommendation using an APC-conjugated antibody (clone PCH 101) or an APC-conjugated isotype control (rat IgG2A). Flow cytometry analysis was performed on the LSR II flow cytometer (BD Biosciences) using BD FacsDiva software (BD Biosciences). Flow-Jo software (Treestar, Ashland, OR) was used solely for graphic presentation in this publication. A minimum of 250 000 total cells was acquired. Lymphocytes were gated by forward-sideward scatter, followed by gates for CD3⁺ and subsequently for CD4⁺ cells. CD25⁺ and FOXP3⁺ cells were analyzed and described as their respective subpopulations within CD4⁺ cells.

Peripheral mononuclear cells (PBMCs) were separated using Ficoll-Hypaque density gradient centrifugation (Organon Teknika, Durham, NC) and subsequently frozen in RPMI 1640 complete medium (CM; Life Technologies, Gaithersburg, MD) supplemented with 20% heat-inactivated fetal calf serum (FCS) and 10% dimethyl sulfoxide (DMSO) according to standard protocols. Before use, frozen cells were thawed, washed, and suspended in RPMI 1640 (Mediatech, Herndon, VA) supplemented with Hepes buffer, gentamicin, and 10% pooled AB serum (Sigma Chemical, St Louis, MO). CD4⁺ cell populations were purified with immunomagnetic beads (Dynal Biotech, Oslo, Norway), which were detached from isolated cells by using DetachaBead (Dynal Biotech).

RNA isolation was performed using RNeasy mini kits (Qiagen, Valencia, CA). Total RNA was eluted with water and stored at −80°C. For reverse transcription of mRNA and cDNA synthesis, 1 μg total RNA was reverse transcribed and stored at −20°C until PCR analysis.

Gene expression was measured using an ABI Prism 7900 Sequence Detection System (Applied Biosystems, Foster City, CA) as described previously.^37,38  Threshold cycle (CT) during the exponential phase of amplification was determined by real-time monitoring of fluorescent emission after cleavage of sequence-specific probes by nuclease activity of Taq polymerase. The internal control gene for mRNA expression was β-actin. Primer and probe sequences for β-actin were as follows: 5′-GGCACCCAGCACAATGAAG (forward), 5′-GCCGATCCACACGGAGTACT (reverse), FAM-TCAAGATCATTGCTCCTCCTGAGCGC-TAMRA (probe). To detect FOXP3, Assays-on-Demand Gene Expression probes for FOXP3 (Hs 00203958; Applied Biosystems) were used according to the manufacturer's guidelines. To create a standard curve, β-actin and FOXP3 cDNA were amplified by PCR using the same primers designed for qRT-PCR, purified, and quantified by UV spectrophotometry. The number of cDNA copies was calculated by using the molecular weight of each gene amplicon. Serial dilutions of the amplified genes at known concentrations were tested by qRT-PCR. qRT-PCR reactions of cDNA specimens, cDNA standards, and water as negative control (NTC) were conducted in a total volume of 20 μL with TaqMan MasterMix (Applied Biosystems), 400 nM primers, and 200 nM probe. Thermal cycler parameters included 10 minutes at 95°C and 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. Standard curve extrapolation of copy number was performed for both FOXP3 and β-actin. Sample data were normalized by dividing the number of copies of FOXP3 transcripts by the number of copies of β-actin transcripts. All PCR assays were performed in duplicates and reported as mean.

As previously described,²³  a quantitative polymerase chain reaction (PCR)–based analysis of short tandem repeats (STRs) was used to measure donor-recipient chimerism separately in lymphoid and myeloid lineages.

Graphs and statistical analyses were performed with the use of Prism 4.00 for Windows software (GraphPad Software, San Diego, CA). P values less than or equal to .05 were considered significant.

Peripheral blood samples of 16 patients (median age, 65 years) were analyzed for T_regs before transplantation and/or during the first 90 days after transplantation. T_regs were also analyzed in 13 of their donors (median age, 62 years) and in 10 of the SD products. These 16 patients received a median of 1.0 × 10⁸/kg selectively depleted (CD25⁻) T cells and a stem-cell product containing a median of 4.5 × 10⁶/kg CD34⁺ stem cells and a median of 0.25 × 10⁴/kg residual unselected T cells (Figure 1). Acute GvHD occurred in 9 of these patients but was limited to grade I, skin-only in 3 of them. Five patients had grade II aGvHD and one developed grade IV aGvHD of the gut. Six patients developed limited and one extensive chronic GvHD (cGvHD). Disease relapse occurred in 6 patients. Median follow-up time was 243 (range, 67-1624) days. Patient characteristics are displayed in Table 1.

Absolute lymphocyte recovery was studied in patients when both absolute lymphocyte counts (ALCs) and complete lymphocyte subsets were available (Figure 2A-E). The fraction of donor chimerism in selected CD3⁺ T cells was solely studied in patients assessable for lymphocyte recovery (Figure 2F). Absolute lymphocyte recovery was prompt, reaching a median of .632 × 10⁹/L (632 cells/μL) (range, .215-2.883 × 10⁹/L [215-2883 cells/μL]) 30 days after transplantation compared with patients before transplantation (median, 1.106 × 10⁹/L [1016]; range, 0.14-4.450 × 10⁹/L [140-4450 cells/μL]) and donors (median, 1.578 × 10⁹/L [1578 cells/μL]; range, .957-2.217 × 10⁹/L [957-2217 cells/μL]) (Figure 2A). Based on CD3 surface expression, absolute lymphocyte counts (ALCs) were split into a subset of CD3⁺ T cells and a subset of CD3⁻ cells consisting predominantly of natural killer (NK) cells since B cells recover relatively late after transplantation. Absolute T-cell counts reached from 92 to 1212 (median, 347) cells/μL at day 30 and increased slowly to 84 to 1456 (median, 397) cells/μL at day 90 but remained below the median values for donors and patients before transplantation (Figure 2B). The CD4⁺ (Figure 2D) and CD8⁺ (Figure 2E) subsets showed the same trend but not CD3⁻ cells (Figure 2C), where median cell numbers 30, 60, and 90 days after transplantation exceeded pretransplantation levels and approached those of the donors. Median donor T-cell chimerism was 97% at day 30, 94% at day 60, and 91% at day 90 (Figure 2F).

T_regs were measured either by flow cytometry or by qRT-PCR. Figure 3 displays the results of flow cytometry for CD25 and FOXP3 and the qRT-PCR analysis for FOXP3 mRNA in CD4⁺ T-cell populations. In the representative patient (UPN 301), all subsets of samples were available for the donor, the CD25-depleted SD product, and the patient before and 30, 60, and 90 days after transplantation. Donor T-cell chimerism was already 97% at day 30 after transplantation and slightly decreased to 94% at day 60 and to 91% at day 90. This patient did not develop aGvHD. The donor cells contained 2.3% CD25⁺ FOXP3⁺ cells (Figure 3A). As expected, the SD product was devoid of CD25⁺ FOXP3⁺ cells (Figure 3A). This patient had 4.0% CD25⁺ FOXP3⁺ cells before transplantation and recovered as early as day 30 1.5% CD25⁺ FOXP3⁺ cells increasing to 4.6% and 5.3% at days 60 and 90 (Figure 3A). With regard to all CD4⁺ FOXP3⁺ cells, this posttransplantation trend could be observed by both flow cytometry (Figure 3B) and by qRT-PCR (Figure 3C). The SD product contained CD4⁺ FOXP3⁺ cells despite complete CD25 depletion (Figure 3A-C). Figure 3A shows that 2.9% of the CD4⁺ cells in the SD product were CD25⁻ and FOXP3⁺, and Figure 3C shows that FOXP3 mRNA-containing CD4⁺ cells persisted after CD25 depletion. CD25⁻ FOXP3⁺ CD4⁺ cells could be also found in the donor (3.8%) and in the patient before (2.9%) and 30 (1.9%), 60 (2.6%), and 90 (2.4%) days after transplantation (Figure 3A). In this patient, the fraction of CD25⁺ FOXP3⁻ T_effs declined from 4.9% at day 30 to 1.7% at day 90. After transplantation, the majority of CD4⁺ T cells expressed CD27. In CD4⁺ cells, expression of CD27 in both CD25⁻ and CD25⁺ T_regs was relatively increased compared with FOXP3⁻ T cells (data not shown).

T_reg recovery was confirmed in the entire study population (displayed in Figure 4). Median FOXP3 mRNA levels increased after transplantation and exceeded the levels of both the donors and the patients before transplantation (Figure 4A). The increase in FOXP3 mRNA levels was associated with a relative increase of FOXP3⁺ and CD25⁺ cells within CD4⁺ cells as depicted in Figure 4B-C. In contrast, CD25⁺ FOXP3⁻ T_effs levels declined after transplantation with the lowest median values observed 90 days after transplantation (Figure 4D), whereas median levels of CD25⁺ FOXP3⁺ T_regs increased after transplantation reaching their highest values 90 days after transplantation (Figure 4E). All SD products contained FOXP3 mRNA and FOXP3 protein but were completely negative for CD25 (Figure 4A-C). The FOXP3 content in the SD product was attributed to a CD25⁻ FOXP3⁺ fraction of T_regs that persisted after CD25 depletion. These CD25⁻ T_regs were also present in patients before and after transplantation and at lower levels in donors (Figure 4F).

Absolute numbers of T_regs were calculated for peripheral blood samples of patients before and after transplantation as well as of their respective donors for whom a complete set of ALCs and flow cytometric profiles were available (Figure 5). Both the medians of absolute FOXP3 copy numbers per cell per volume and the absolute number of CD4⁺ FOXP3⁺ cells per volume increased during the first 90 days after transplantation coinciding with the results described for relative T_reg recovery (Figure 5A,B). Thirty days after transplantation, patients had a median of 10 CD4⁺ FOXP3⁺ cells/μL, increasing to 14 at day 60 and to 19 at day 90. Patients before transplantation had a median of 18 and donors of 16 CD4⁺ FOXP3⁺ cells/μL (Figure 5A,B). Median absolute numbers of all CD25⁺ CD4⁺ cells (Figure 5C) and CD25⁺ CD4⁺ FOXP3⁺ T_regs (Figure 5E) followed this trend but not median absolute numbers for CD25⁺ CD4⁺ FOXP3⁻ T_effs (Figure 5D) and CD25⁻ CD4⁺ FOXP3⁺ T_regs (Figure 5F).

To explore the association of T_regs with the onset of aGvHD, we compared the levels of FOXP3⁺ CD4⁺ T cells between patients with no or grade I aGvHD and patients with aGvHD (≥ grade II). We investigated the effects of T_regs in the patient before and 30 days after transplantation, the donors, and the SD products at the onset of aGvHD (≥ grade II) (Figure 6). Levels of FOXP3⁺ cells in the CD4⁺ content in patients before and early after transplantation and in the SD product were not associated with the onset of aGvHD (Figure 6A-C). However, aGvHD (≥ grade II) was restricted to 5 patients whose donors had significantly (P = .019) fewer T_regs compared with those with no or grade I aGvHD (Figure 6D). This effect of the donor T_regs was also seen in association with the absolute number of CD25⁺ CD4⁺ FOXP3⁺ T_regs (P = .045) but not with CD25⁻ CD4⁺ FOXP3⁺ T_regs (P = .22) (Figure 6E,F). There was no association of T_regs with cGvHD or relapse (data not shown).

The stimulation of donor T cells with recipient antigen-presenting cells (APCs) and the concurrent removal of host-reacting T cells during the selective depletion process promises transplantation outcomes with a reduced incidence of aGvHD, while providing a fully functional donor immune system retaining its antileukemic potential uninhibited by immunosuppression. We previously showed that removal of host-reactive donor T cells from allografts by anti-CD25 IT was clinically feasible and reduced the frequency of severe aGvHD in a high-risk group of elderly patients undergoing matched-sibling transplantation who received only low-dose cyclosporine for immunosuppression.²³  However, it was a concern that acute and chronic GvHD in our patients could have been associated with the concurrent removal of T_regs with the alloactivated CD25⁺ T cells. In the present study, we found efficient T_reg reconstitution in all patients. Using FOXP3 as a marker to distinguish between effector and regulator populations in CD4⁺ cells, we found a relative decline of T_effs in contrast to a relative increase of T_regs over different time points after transplantation diminishing the effector-to-regulator ratio in favor of a more suppressive T-cell environment.

To further characterize T_regs, we also investigated the expression of CD27, a TNFR-family member that was found to be consistently expressed on T_regs.³⁹  CD27 is absent on effector T cells and is rapidly lost on CD27⁺ naive and memory T cells after activation.³⁹  In our series, the majority of reconstituted CD4⁺ T cells were CD27⁺. Nevertheless, CD27 expression in both CD25⁺ and CD25⁻ T_regs was relatively increased compared with FOXP3⁻ CD4⁺ T cells.

Regarding potential sources for T_reg reconstitution, we considered the possibility that the reduced-intensity conditioning regimen could have conserved some patient-derived T cells leading to mixed T-cell chimerism after transplantation. However, by chimerism analysis the T-cell repertoire of our patients was found to be predominantly of donor origin. Furthermore, 5 patients who had full donor T-cell chimerism also reconstituted considerable numbers of T_regs as early as 30 days after transplantation. Therefore, it is unlikely that the T_regs reconstituting in our patients were derived from recipient T cells surviving the conditioning procedure. In view of the advanced age of our patient population and the fact that we studied early (fewer than 100 days after transplantation) T_reg reconstitution, the potential thymic contribution in the de novo generation of T_regs from CD34⁺ stem cells can be disregarded.⁴⁰  Thus, the donor T cells appear to be the most likely source of T_reg reconstitution. Donor T_regs could have been derived from 2 sources, a predominant population of CD25-depleted T cells in the SD product (1 × 10⁸/kg) and a 4-log smaller amount of residual T cells (0.25 × 10⁴/kg) contained in the stem-cell product. Although we confirmed that SD products did not contain CD25⁺ T_regs, we identified a population of CD25⁻ (FOXP3⁺) T_regs persisting after CD25 depletion. FOXP3, the recently identified key player in T_regs, is predominantly but not exclusively expressed in CD4⁺ CD25⁺ T cells.^33-35  FOXP3 expression in CD25⁻ CD4⁺ cells also exerts suppressive and regulatory function as recently shown in mice⁴¹  and humans.⁴²  We also found these CD25⁻ T_regs in our patients before and after transplantation and in the donors, suggesting that the residual unselected T cells in the stem-cell product provide a small number of CD25⁻ and CD25⁺ T_regs. Thus, it seems likely that T_reg recovery following SD is derived mainly from CD25⁻ T_regs which escape the depletion process with a further contribution of CD25⁻ and CD25⁺ T_regs delivered in the stem-cell product. Regarding the developmental pathways of T_regs, it has already been suggested that CD25⁻ T_regs may up-regulate their CD25 receptor following antigen stimulation⁴³  and thereby contribute to the pool of CD25⁺ T_regs. This conversion would explain why we observed increasing levels of CD25⁺ T_regs after transplantation, while levels of CD25⁻ T_regs remained rather stable. Finally, all these CD25⁺ donor-derived T_reg fractions could have undergone lymphopenia-driven expansion⁴⁴  after transplantation, thereby contributing to GvHD control and compensating for thymic failure of de novo T_reg production.

Reconstituting donor postthymic T cells are responsible for establishing early immune function after SCT. While early studies investigating the relationship between T_reg recovery and occurrence of aGvHD in humans were limited by the difficulty in distinguishing T_reg from T_eff populations, FOXP3 analysis has provided a more consistent picture where high FOXP3 levels in blood or tissue were associated with less aGvHD^24,25,27,32  Here, we found an association between the donor T_reg content and the occurrence of clinically significant aGvHD. In contrast, neither the reconstituted level of T_regs in the patient nor the T_reg content of the SD product was associated with GvHD suppression. These results concur with our findings in unselected, T-cell–depleted SCT and emphasize that donor-derived T_regs contribute to GvHD suppression in the host at a very early stage.²⁴ 

In mice, CD25 depletion was associated with the development of significant autoimmune disease,⁴⁵  and in humans a clinical trial attempting the in vivo CD25 depletion after allografting was terminated due to an increased rate of aGvHD,⁴⁶  attributable to T_reg removal. In this study, we showed efficient T_reg reconstitution despite giving a CD25-depleted allograft. Thus, the occurrence of aGvHD in our series of SD stem-cell transplant recipients cannot simply be explained by the removal of T_regs, but rather to an insufficient removal of alloreactive T cells by the SD process. Indeed, we previously reported that in patients developing aGvHD involving the gastrointestinal tract and the liver, there was persistent donor-versus-recipient alloreactivity as measured by helper T lymphocyte precursor frequencies.²³  Either the recipient T-cell APCs used to stimulate the donor did not present all the potential GvHD-related antigens or the subsequent depletion did not remove all the alloactivated donor cells. The latter scenario could be partly due to a down-regulation of CD25 antigen during the coculture period allowing some alloactivated cells to escape the depletion process. Thus, future clinical developments of SD may also need to focus on varying the type of recipient APCs (eg, Epstein-Barr virus–transformed lymphoblastoid cell lines²¹  or unmanipulated PBMCs²² ) as well as explore novel targets of donor T-cell activation (eg, CD137¹⁴ ) or different depletion techniques (eg, TH9402-based photodepletion^47,48 ).

In this study, we showed for the first time that SD using CD25 as the depleting target may protect against aGvHD, both by allowing efficient T_reg reconstitution as well as by allodepletion. It should be emphasized that our study population is small and these results will need validation in a larger patient cohort. The SD approach nevertheless seems worthwhile since the technique offers the possibility of removing GvHD reactivity while preserving other donor immune function. Unlike other techniques involving T-cell subset depletion, SD appears to least perturb the balance of immune reactivity between regulator and effector cells, and conservation of T_reg recovery may provide further GvHD protection when allodepletion is incomplete.

S.M. was supported in part by a grant of the Dr-Mildred-Scheel-Stiftung for Krebshilfe, Germany.

National Institutes of Health

Contribution: S.M. conceived and designed the study, performed experiments, analyzed data, performed statistical analyses, and wrote the paper; K.R. conceived and designed the study, performed experiments, analyzed data, and commented on the paper; B.N.S. collected patient data and provided clinical care; R.N. performed experiments; A.S.M.Y. advised on experimental design and commented on the paper; J.S. was involved in development and chemical manufacture of the CD25-immunotoxin for selective depletion and commented on the paper; R.K. provided the chimerism data; V.G. was involved in development and chemical manufacture of the CD25-immunotoxin for selective depletion; E.J.R. supervised the clinical selective depletion process, provided data, and commented on the paper; S.R.S. was the principal investigator on the selective depletion trial, collected patient data, and provided clinical care; E.S.V. was involved in development and chemical manufacture of the CD25-immunotoxin for selective depletion and commented on the paper; A.J.B. supervised this study, supervised the selective depletion trial, provided clinical care, and wrote the paper. S.M. and K.R. contributed equally to this work.

Conflict-of-interest disclosure: The authors declare no competing financial interests.

Correspondence: Stephan Mielke, Stem Cell Allogeneic Transplant Section, Hematology Branch, NHLBI, NIH, Bldg 10 CRC Rm 3-5288, 10 Center Dr, MSC 1202, Bethesda, MD 20892-1202; e-mail: mielkes@nhlbi.nih.gov.