Early T Cell Activation Metrics Predict Graft-versus-Host Disease in a Humanized Mouse Model of Hematopoietic Stem Cell Transplantation

Graft-versus-host disease (GVHD) is a major complication following allogeneic hematopoietic stem cell transplantation (HSCT) that is associated with nonrelapse mortality, delayed immune reconstitution, elevated risk of infections, and overall poor quality of life (1, 2). The central feature of GVHD is the reactivity of donor T cells against host tissues, primarily manifesting as liver, gut, and mucous membrane pathology (3). Despite the risk of GVHD after allogeneic HSCT, the probability of any given patient developing GVHD is difficult to predict because it is dependent on a variety of factors, including the number of major and minor MHC mismatches between donor and recipient, the type of pretransplant conditioning regimen (myeloablative versus reduced intensity versus nonmyeloablative), type of graft source used for the transplant (bone marrow [BM], G-CSF mobilized peripheral blood [MB], or umbilical cord blood [CB]) and the nature of the posttransplant GVHD prophylaxis (traditional calcineurin regimens versus posttransplant Cytoxan [cyclophosphamide]/anti-thymocyte globulin) (1, 2). Without a clear consensus on optimal treatment regimens, murine model systems have been integral in elucidating the biology of GVHD development and pathology.

Studies in murine models of GVHD have revealed a general mechanism of immunological events prior to disease pathology. In brief, host hematopoietic and nonhematopoietic APCs that are not cleared by the conditioning regimen become activated in the presence of damage-associated molecular patterns released during the conditioning (3–7). The resulting inflammatory host APCs then activate donor naive T cells transplanted within the graft against host Ags, which leads to TH₁, TH₁₇ and CD8⁺ cytotoxic responses against host organs (3, 4, 8).

Whereas murine models have provided a critical mechanistic understanding of processes leading up to GVHD, these models are limited in their capacity to directly evaluate the impact and functional differences associated with the various clinically used graft tissues. Xenotransplantation models using immunodeficient mice provide an opportunity to address these gaps in our understanding of the posttransplant behavior of different sources of human graft tissues. However, possibly as a result of inefficient competition for murine hematopoietic factors, pretransplant irradiation is required for human cells to successfully engraft most strains of immunodeficient mice (9–11). Because irradiation causes damage that promotes GVHD by activating host responses, we sought an approach that would allow us to investigate the inherent propensity of human T cells to cause pathology in a host environment lacking radiation-associated damage. To do this, we used a recently derived immunodeficient mouse strain (NBSGW) that is highly permissive for transplanted human immune cells in the absence of conditioning (12). This novel model revealed distinct differences among the various human hematopoietic graft tissues in the cellular dose required to cause GVHD and identified early T cell activation metrics that are predictive of subsequent GVHD pathology, irrespective of the source of hematopoietic tissue used.

Human BM cells and G-CSF MB were collected from the remnants left over in de-identified bags and filters used for clinical HSCT procedures. De-identified human CB samples were acquired from the University of Colorado ClinImmune Labs CB bank or the Medical College of Wisconsin’s tissue bank. Venous peripheral blood (PB) was drawn from healthy consenting donors. Graft sources were isolated by Ficoll density gradient centrifugation (1100 × g for 15 min with 0 brake) to remove RBCs, neutrophils, and other nonleukocytes. Where indicated, STEMCELL Technologies RosetteSep T Cell Enrichment Cocktail was used (catalog no. 15061) to remove lineage-positive cells (excluding T cells), leaving untouched CD34⁺ hematopoietic stem and progenitor cells (HSPCs) and T cells; T cells were depleted using STEMCELL Technologies RosetteSep T cell Depletion Kit (catalog no. 15661). HSPCs were isolated using STEMCELL Technologies EasySep Human CD34 Positive Selection Kit II (catalog no. 17856), which first depletes lineage-positive cells, followed by magnetic labeling of CD34⁺ cells and isolation using a magnetic column. Cells were counted, washed, and resuspended in PBS prior to injection into mice.

The immunodeficient mouse strain NBSGW (NOD.Cg-Kit^W-JTyr⁺Prkdc^scidIL2rg^tm1Wjl/ThomJ) was purchased from The Jackson Laboratory and bred and maintained in specific pathogen–free facilities using aseptic housing. Equal numbers of male and female mice between 6 and 10 wk of age were used for all experiments. Human cells were suspended in 150 μl of PBS and injected retro-orbitally. Mice were weighed and monitored weekly for visible signs of GVHD, with scoring carried out as follows: 0 = no signs of GVHD, 1 = no weight loss but visible signs of GVHD (hunching, lethargy, ruffled fur), 2 = 0–5% weight loss, 3 = 5–10% weight loss, 4 = >10% weight loss. Mice that lost 10% or more of their maximum body weight were euthanized. Blood was drawn at the indicated time points, and after 3 mo all mice were euthanized, and the spleen, one lobe of the liver, and the left leg were collected for further processing. For some experiments, LPS was coinjected with human cells at 1 ng/ml or i.p. injected at 50 ng/g (Invitrogen).

Single-cell suspensions were blocked in 10% human serum and stained for flow cytometric analysis using the following fluorophore-conjugated Abs purchased from BioLegend: CD3 (OKT3), CD4 (SK3), CD8 (SK1), CD10 (HI10A), CD19 (HIB19), CD28 (C28.2), CD33 (HIM3-4), CD34 (8G12), CD38 (HIT2), mCD45.1 (A20), CD45RA (HI100), CD45RO (UCHL1), CD56 (HCD56), CD66b (610F5), CD123 (6H6), IFN-γ (4S.B3), TNF-α (MAb11), OX40 (Ber-ACT35), ICOS (C398.4A), CTLA4 (BNI3), PD1 (EH12.2H7), and PanHLA (W6/32). Stained samples were resuspended in PBS and analyzed on a BD LSR II flow cytometer equipped with three lasers (nine filter channels) and quantified using Precision Count Beads from BioLegend. Data analysis was performed using FlowJo V10.5.

Cultures of BM-HSPCs were performed using STEMCELL Technologies StemSpan Media (catalog no. 09650) supplemented with StemSpan CD34⁺ Expansion Supplement (catalog no. 02691) for 7 d at 37°C and 5% CO₂. BM-HSPCs were stained prior to culturing with Thermo Fisher Scientific’s CellTrace Violet (CTV; catalog no. C34557). Cell culture supernatants or mouse plasma was diluted either 1:5 or 1:25 in ELISA Buffer, respectively, before performing an IFN-γ–specific ELISA using the Ab clones MD-1 (coating), biotinylated 4S.B3 (detection), and a streptavidin-HRP Ab (BioLegend).

Graphs and statistical tests were completed using GraphPad Prism 6. Means are shown for data plotted on linear axes, whereas geometric means are used when the data are presented on a logarithmic axis. Trendlines on plots of logarithmic data show results of nonlinear regression analyses. Correlations were determined using either a parametric Pearson coefficient (for data following a Gaussian distribution) or a nonparametric Spearman coefficient and denoted as a correlation coefficient (r). Unpaired, nonparametric t tests were used to assess significance of aggregated data shown on columnar scatter plots. Receiver operator characteristic (ROC) analyses were done by GraphPad Prism, based on mice having no GVHD as the control condition. For the T cell blasting analysis, mice with <10 collected T cell events were excluded from the analysis to avoid skewing the result.

All work involving human cells and tissues was performed in accordance with Institutional Review Board protocol 2018-0304 (J.E.G.), 2016-0298 (P.H.), and 2017-0870 (J.E.G.). All animal work was performed in accordance with University of Wisconsin-Madison Institutional Animal Care and Use Committee–approved protocol number M005199.

Immunodeficient NBSGW mice have been shown to accept purified human HSPC without prior conditioning and no evidence of GVHD development (12, 13). However, clinical HSCT protocols are typically performed using unmanipulated graft sources that contain a mixture of cell types. To investigate the ability of clinical human grafts to mediate GVHD in a nonconditioned environment, we injected total mononuclear cell (MNC) BM cells (BM-MNC) retro-orbitally into NBSGW mice at various doses and monitored them for 13 wk (Fig. 1A). There was a dose-dependent increase in GVHD mortality (>10% loss of maximum body weight) and GVHD score (based on weight loss, hunching, squinting, and lethargy) with a dose of 1 × 10⁷ BM-MNC yielding an ∼50% penetrance of lethal GVHD (Fig. 1B, 1C). Interestingly, analysis of the BM of mice with or without GVHD at the time of euthanasia revealed no statistical difference in the total number of human T cells in the murine BM or spleen (Fig. 1D, 1E) but did show a distinct difference in the number of other human immune cell types (Fig. 1F). Moreover, the number of other human immune cells (i.e., human cells excluding T cells) in the murine BM showed a significant negative correlation with GVHD severity (Fig. 1G). Thus, despite not inducing damage-associated ligands in this model, GVHD still occurred and was not associated with a significant increase in the absolute numbers of human T cells but instead, was negatively associated with the expansion of non-T cell human immune lineages.

Importantly, however, we found that GVHD was not an inevitable outcome of human xenotransplantation in this system. The percentage of mice developing GVHD was dependent on the dose of human BM cells administered (Fig. 1B). Mice that received up to 4 × 10⁶ human BM cells had human chimerism that included T cells but did not develop detectable GVHD within 3 mo (Fig. 1B, 1D). Moreover, we found that mice that received 1 × 10⁷ human BM cells had a 50% chance of developing lethal GVHD that was consistent across multiple human BM donors (Fig. 1C). Additionally, administration of a sublethal dose of LPS at days 0 and 3 posttransplant produced a trend toward increased incidence of GVHD in mice that received 1 × 10⁷ human BM cells (Fig. 1H, 1I). These results demonstrated that the induction of GVHD is not an intrinsic feature of xenotransplantation and, instead, is related to human cell dose and probably also to the presence of inflammatory factors after transplantation.

GVHD pathology in the NBSGW mice was characterized by widespread lymphocytic infiltration of the liver, kidney, lung, and salivary gland, whereas the skin and gastrointestinal tract remained clear (Fig. 2A). Liver tissue collected at the time of euthanasia and scored for lymphocytic infiltration by blinded assessment of H&E-stained sections revealed a dose-dependent increase in liver pathology (Fig. 2A, 2B). To further show GVHD mice developed liver damage, we assayed the plasma of GVHD and non-GVHD mice for the liver enzymes ALT, ALP, and AST and compared them to a murine reference range (14). Most mice with GVHD had elevated levels of AST and some also showed elevated ALT, whereas levels of ALP, blood urea nitrogen, and albumin were mostly unaffected (Fig. 2C, data not shown). Lastly, all mice that developed GVHD had marked decreases in body weight, and that was the primary phenotypic indicator of GVHD pathology used in this study (Fig. 2D, 2E).

Studies in murine models indicate that donor T cells but not donor APCs are required for the induction of GVHD (5). We investigated the role of donor T cells and donor APCs in our model by transplanting total BM-MNCs compared with either T cell–depleted BM or isolated BM-T cells (lineage-negative CD34⁺ HSPCs were included as an internal control, see 2Materials and Methods). Mice receiving T cell–depleted grafts showed no evidence of GVHD, whereas mice given isolated BM–T cells developed GVHD at the same frequency as BM-MNC when controlling for injection dose (Fig. 3A–D). Mice given BM-MNC and isolated BM–T cells also had an equivalent number of T cells and human non-T cells derived from the cotransplanted HSPCs in their BM and spleen at the time of euthanasia (Fig. 3E, 3F). These data show that, similar to what has been observed about the induction of GVHD in murine models, transplantation of human T cells but not cotransplantation of human APCs is required for GVHD in this model.

To investigate the changes in the human T cell population over time, mice transplanted with either BM-MNC or isolated BM–T cells were bled at regular intervals, and the human T cells in the blood were analyzed and quantified by flow cytometry. Mice that eventually developed GVHD had significantly higher numbers of human T cells in the blood by 3 wk posttransplant (Fig. 4A, Supplemental Fig. 1A). The difference in blood T cell numbers remained significant throughout the remainder of the experiment (Fig. 4A), but differences in T cell numbers were less apparent in the BM and spleen of mice given BM-MNC when compared between one BM-MNC dose (Fig. 1E) or multiple doses (Fig. 4B). However, mice given isolated BM-T cells that went on to develop GVHD showed significantly elevated T cell numbers in BM, spleen, and PB throughout the experiment (Supplemental Fig. 1A, 1B). T cell expression of ICOS, CTLA-4, and PD-1 were upregulated compared with pretransplant controls but did not differ between mice that ultimately did or did not develop GVHD (Fig. 4C, Supplemental Fig. 2A). Furthermore, almost all T cells transitioned to CD45RO⁺ by 6 wk posttransplant in both GVHD and non-GVHD mice, suggesting that T cells gained antigenic experience regardless of whether GVHD pathology developed (Fig. 4G, Supplemental Figs. 1C, 2B). Thus, GVHD may not simply be an outcome of antigenic stimulation but may instead depend on the nature of the T cell activation process.

To investigate this further, we examined the frequency of blasting T cells (an increase in cell size that is classically associated with rapid lymphocyte proliferation) in the blood of BM-MNC and isolated BM–T cell–transplanted mice over the course of the 13 wk experiment. Interestingly, there was a significant difference in the percentage of blasting T cells between GVHD and non-GVHD mice as early as 1 wk posttransplant, and that difference was maintained in the BM and spleen (Fig. 4D–F, Supplemental Fig. 1D, 1E). To assess whether blasting frequency is correlated with T cell functional status, we took splenic cells from BM-MNC mice collected at the time of euthanasia and incubated them in vitro in the presence or absence of PMA/ionomycin before assessing their intracellular IFN-γ, IL-17A, and TNF-α production. There was no difference in the number of T cells producing either IFN-γ, IL-17A, or TNF-α after PMA/ionomycin treatment (Fig. 4H–J, Supplemental Fig. 2C). However, in the absence of PMA/ionomycin treatment, mice that had a higher frequency of blasting T cells and went on to develop GVHD showed a significantly elevated frequency of T cells that stained positively for IFN-γ and IL-17A (Fig. 4H, 4I). In contrast, TNF-α production by nonstimulated T cells did not differ significantly (Fig. 4J). These results suggested that T cells in the non-GVHD mice did not have an inherent defect in their activation potential but instead had not been activated in vivo to produce IFN-γ or IL-17A, whereas T cells from mice that went on to develop GVHD were likely secreting IFN-γ and IL-17A at early time points in vivo.

Therefore, we next investigated the role of IFN-γ within our model. Plasma taken from regular blood draws was analyzed for IFN-γ by ELISA. We observed a significant difference between the GVHD and non-GVHD mice as early as 6 wk posttransplant in the BM-MNC condition and 3 wk in the isolated BM–T cells condition (Fig. 5A, 5B). As noted earlier (Fig. 1F, 1G), mice with GVHD had reduced numbers of human non-T cell lineages in their BM. To investigate whether IFN-γ might be involved in this effect, we collected the supernatant from murine BM harvests and analyzed them for IFN-γ by ELISA. Samples from mice with GVHD showed significantly higher amounts of IFN-γ that were negatively correlated with the presence of human non-T cell lineages in the murine BM (Fig. 5C, 5D).

It is well established in murine model systems that chronic IFN-γ exposure can prevent HSPC proliferation (15–17). Therefore, we investigated whether IFN-γ had a similar effect on HSPCs isolated from our human BM samples. Isolated human HSPCs were cultured for 7 d alone or in a transwell culture with autologous BM–T cells or CD3/CD28–activated BM-T cells with their proliferation monitored with a CTV dye and overall cell number quantified. Although there were no differences between HSPCs cultured alone or in the presence of autologous T cells, there was a significant reduction in proliferation and cell number of HSPCs cultured in the presence of activated T cells (Fig. 5E, 5F). We also confirmed that IFN-γ was produced only in the condition with activated T cells and that the HSPCs had detectable levels of IFN-γR1 (CD119) (Supplemental Fig. 3). Lastly, isolated HSPCs were given rIFN-γ at various doses and showed a significant reduction in cell number in the presence of as little as 1 ng/ml IFN-γ (Fig. 5G).

Currently, there is no clear consensus on the optimal source of tissue for allogeneic HSCT. The most commonly used tissues are G-CSF MB, BM, and CB (in that order), but their use also varies among clinics (1, 2, 18). Within the clinic, CB transplantation has been shown to have the lowest rates of acute GVHD, whereas MB in the context of allogeneic transplants have the highest rates of acute GVHD (1, 2). Despite these observations, there have been few studies that have directly investigated the propensity of each graft tissue to mediate GVHD in a controlled laboratory setting. To directly compare the propensity of these three types of tissue to produce GVHD in our model, we transplanted nonconditioned NBSGW mice with titrated cell doses and monitored them for signs of GVHD over time. Similar to what has been observed clinically, CB had the lowest propensity to produce GVHD requiring ∼7.3 × 10⁷ CB-MNC to produce lethal GVHD in 50% of the mice (Fig. 6B, 6E). Surprisingly, however, BM produced GVHD at lower cell doses than MB, with calculated LD₅₀ values of 1 × 10⁷ and 2 × 10⁷ MNC, respectively, which is not explained by differences in the cellular composition of the grafts (Fig. 6A, 6C, 6E, Supplemental Fig. 4). To further investigate, we compared transplantation of PB from healthy control subjects to transplantation of MB from G-CSF–treated donors. Strikingly, MB had a 10× higher LD₅₀ than PB (Fig. 6C–E). Hence, G-CSF treatment of donors may result in reduced propensity of the graft tissue to produce GVHD, compared with normal PB (19). Notably, however, for the mice that did get GVHD, the time to death was shortest for mice that received MB tissue (Fig. 6F). Thus, this model suggests that MB has a comparatively low propensity to produce GVHD, but when pathology does occur, it may be more severe than that produced by other types of tissue.

We next assessed whether the T cell activation metrics we identified as early events that precede the onset of GVHD pathology in mice that received BM transplants (Figs. 4, 5) show a similar predictive value for other types of graft tissue. Regardless of the type of human tissue transplanted, mice that went on to develop GVHD had an elevated frequency of blasting T cells at week 1 (Fig. 7A), a higher burden of T cells in the blood at week 3 (Fig. 7B), and an increased concentration of IFN-γ in their plasma at week 6 (Fig. 7C). We analyzed each parameter using an ROC curve that tests the efficacy of each metric to predict an outcome. This analysis revealed that each of the three parameters was significantly predictive of GVHD pathology between 1 and 6 wk posttransplant across all four graft sources tested, with the percentage of blasting T cells in blood showing the most accuracy at the earliest time point (Fig. 7D–F). These results establish that a proliferative burst by circulating T cells as early as 1 wk posttransplant is a reliable predictor for subsequent development of GVHD pathology regardless of the type of human tissue transplanted and even in the absence of radiation-induced damage in the host.

In this study, we have established a nonconditioned immunodeficient mouse model to evaluate the GVHD propensity of clinically used human graft sources and defined several early T cell activation metrics that are predictive of GVHD pathology irrespective of graft source. Although using immunodeficient mice provides a powerful approach to study in vivo human immunological events, there are important limitations to consider with xenogeneic transplant models (20). Cytokines that are critical for lymphocyte homeostasis, such as IL-2 and IL-7, and cytokines that are required for HSPC function, such as stem cell factor and stromal cell–derived factor 1, are all fully cross-reactive between mice and humans (12). However, the full extent of cross-reactive cytokines between mice and humans is not yet known, and many key cytokines (such as IL-3, IL-6, IL-15, M-CSF, GM-CSF, and IFN-γ) are not highly cross-reactive (12, 21). Additionally, the level of stimulation received by human T cells from murine MHC and costimulatory molecules remains unclear (22). Our data show that, whereas pathological activation of human T cells is not an inevitable outcome following xenotransplantation, the murine environment is capable of activating human T cells to produce GVHD pathology regardless of whether human APCs are cotransplanted (Figs. 1–3).

Another unique aspect of this study is the use of nonconditioned mice. Whereas harsh myeloablative conditioning regimens have classically been used to eliminate malignant cells, conditioning regimens with lower toxicity are now used on older patients and those with specific comorbidities (23). The trend of using reduced toxicity conditioning regimens has continued with the introduction of a bead-based conditioning protocol, which has recently entered clinical trials, that allows for engraftment without causing any damage to the host (24–26). Because these types of conditioning protocols may become increasingly common in HSCT protocols in the future, it is imperative to understand the potential of HSCT grafts to mediate GVHD in a noninflammatory environment.

Acute GVHD in the clinic is characterized by donor T cell–driven pathologies of the gastrointestinal tract, liver, and skin (3). We confirmed that our model recapitulates the infiltration of lymphocytes into the liver and that the resulting damage is measurable by the release of the liver enzymes AST/ALT (Fig. 2B, 2C). Interestingly, we did not detect any infiltration or damage in the murine gastrointestinal tract or skin (Fig. 2A). Whereas gastrointestinal GVHD is extremely common in clinical cases, it is also an organ that is predominantly damaged by conditioning regimens (27, 28). Thus, the absence of a conditioning regimen in our model may prevent the induction of gastrointestinal GVHD, although additional experiments are required to confirm this.

Another aspect of clinical HSCT that our model recapitulates is the inverse correlation between systemic IFN-γ concentration and de novo expansion of the human hematopoietic compartment in the BM of our mice (Fig. 5). It has become well established from both murine studies and immune reconstitution data in the clinic that chronic IFN-γ exposure severely limits the expansion and differentiation of HSPCs (15–17, 29). Clinically, this often manifests as a drop in neutrophil and platelet counts after the onset of GVHD pathology. In our model, the total human non-T cell compartment was reduced in GVHD mice (Fig. 1F, 1G), which is due to the adverse effect of GVHD-associated cytokines (e.g., IFN-γ) on HSPC function (Fig. 5). The inclusion of human HSPCs within the grafts transplanted into our model system also highlights the possibility of de novo T cell development. Importantly, we and others have not detected any de novo T cell development in the murine BM when we transplanted T cell–depleted or isolated HSPCs (Fig. 3 and data not shown). Furthermore, necropsies of these mice have shown that the murine thymus atrophies shortly after birth, rendering it nonfunctional at the ages used in this study (12, 13).

A key feature of our model system is the incomplete penetrance of GVHD pathology at intermediate cell doses. This suggests that GVHD is not simply a function of transplanting human T cells into a xenogeneic environment and is instead associated with activation events that do not necessarily occur in all similarly transplanted mice (Figs. 1, 6). Whereas the nature of these activation events is currently not clear, we have identified several variables that influence the probability of an individual mouse to develop GVHD (22). We show that cellular dose, graft source, and probably also systemic inflammation, can all modulate GVHD risk. Because our model is highly permissive to transplantation of human cells without prior conditioning, the baseline inflammation within our model is extremely low. Similar to what is reported in the clinical setting, we observed a trend for systemic inflammation (mediated by sublethal doses of LPS) to increase the penetrance of GVHD in our model but found that it is not required for GVHD initiation (Fig. 1F, 1G). In addition to the dose escalation of GVHD penetrance, this data implies that upon transplant, there is a threshold of activation that must occur within the T cell population for later GVHD pathology to develop, which occurs stochastically within each mouse. Thus, both systemic inflammation and increased cellular dose increase the chances of any given mouse of surpassing that activation threshold (30). Importantly, the putative activation threshold appears to be independent of T cell acquisition of antigenic experience, as both T cells from GVHD and non-GVHD mice become CD45RO⁺ and have elevated levels of PD-1, CTLA-4, and ICOS. Thus, the nature of the activation mechanism driving GVHD pathology requires further investigation.

It is well established that clinical transplantation of G-CSF MB is associated with higher rates of GVHD than BM transplantation (1, 2). However, our data suggests that on a per cell basis, BM is more GVHD prone that MB. The most striking difference in GVHD propensity as measured by the calculated LD₅₀ values is the differences between PB and MB. Despite both being comprised of cells circulating in the blood, these two graft sources have an approximate 10-fold difference in LD₅₀ values (Fig. 6). This supports the possibility that G-CSF treatment prior to transplantation may have suppressive effects, although prior studies have come to contradictory conclusions on this point (31–33). These findings highlight the importance of further examination of the biological factors that influence the GVHD propensity of tissues used for HSCT.

Whereas the events leading to T cell activation remain unclear, we have identified three novel biomarkers of early T cell activation that are all predictive of GVHD 3–6 wk before visible pathology and are independent of graft source (Figs. 4, 7). Because of the number of variables impacting the development of GVHD in clinical settings, there is a significant need to identify prognostic indicators of GVHD risk before the onset of pathology. Studies to date have predominantly focused on plasma biomarkers, with ST2 and REG3α both rigorously validated as indicators of GVHD severity, probability of nonrelapse mortality, and steroid-refractory GVHD (34, 35). Additionally, the recovery of microbial diversity or lack thereof after transplant is a biomarker of GVHD development that is currently gaining traction (36). Our data suggest that the percentage of blasting T cells, total T cell burden, and blood IFN-γ concentration may provide important new biomarkers for GVHD risk. Moreover, these parameters may turn out to provide a valuable way to identify risk of subsequent pathological activation for cellular immunotherapies in which T cell activation is involved, such as CAR-T cell therapies. In conclusion, this study outlines a novel model system to investigate differences in human HSCT graft sources as well as the ability to define specific early T cell activation events that are predictive of GVHD pathology.

We acknowledge Melissa Graham and Beth Gray at the University of Wisconsin-Madison Comparative Pathology Laboratory for assistance with histology and Ashley Weichmann at the University of Wisconsin-Madison Small Animal Imaging Facility for assistance with measuring serum analytes.

The authors have no financial conflicts of interest.