Critical role of host T cells in experimental acute graft-versus-host disease
http://www.100md.com
《血液学杂志》
the Departments of Internal Medicine, General Surgery
Pediatrics, University of Michigan Cancer Center, Ann Arbor, MI
the Department of Pathology, University of Florida College of Medicine, Gainesville.
Abstract
T cells localize to target tissues of graft-versus-host disease (GVHD) and therefore we investigated the role of host T cells in the pathogenesis of acute GVHD in several well-characterized allogeneic bone marrow transplantation (BMT) models. Depletion of host T cells in wild-type (wt) B6 recipients by administration of anti-T-cell receptor (TCR) monoclonal antibody reduced GVHD, and T-cell-deficient (-/-) BM transplant recipients experienced markedly improved survival compared with normal controls (63% vs 10%, P < .001). T cells were responsible for this difference because reconstitution of -/- recipients with T cells restored GVHD mortality. -/- recipients showed decreased serum levels of tumor necrosis factor (TNF-), less GVHD histopathologic damage, and reduced donor T-cell expansion. Mechanistic analysis of this phenomenon demonstrated that dendritic cells (DCs) from -/- recipients exhibited less allostimulatory capacity compared to wt DCs after irradiation. Normal DCs derived from BM caused greater allogeneic T-cell proliferation when cocultured with T cells than DCs cocultured with medium alone. This enhancement did not depend on interferon (IFN-), TNF-, or CD40 ligand but did depend on cell-to-cell contact. These data demonstrated that the host T cells exacerbate GVHD by enhancing the allostimulatory capacity of host antigen-presenting cells. (Blood. 2005;106:749-755)
Introduction
Allogeneic bone marrow transplantation (BMT) is a widely performed therapy for many hematologic malignancies. However, acute graft-versus-host disease (GVHD) remains the most important complication of allogeneic BMT. The pathophysiology of GVHD is complex. The target organs of acute GVHD include the gastrointestinal tract, skin, and liver. Several convergent lines of experimental data have demonstrated that donor T cells and host antigen-presenting cells (APCs) are critical for the induction of GVHD.1-3 In addition, a growing body of data also suggest that several other cellular subsets such as host macrophages,4,5 donor natural killer (NK) cells,6,7 and donor NKT cells8 are involved in the pathogenesis of GVHD.
T cells represent a distinct lineage of T cells that evolved early in vertebrate phylogeny in conjunction with T cells and B cells. Antigen recognition by T-cell receptors (TCRs) occurs in a fundamentally different manner from that of T cells. T cells can recognize molecules other than peptides, do not require professional APCs for activation, and use their TCRs to recognize patterns associated with foreign pathogens or tissue injury.9 These cells serve as sentinels to monitor their environment for signs of stress caused by infection, inflammation, and transformation. T cells modulate immune responses by secreting cytokines and by regulating the function of other immune cells. For example, T cells produce either interferon (IFN-) or interleukin 4 (IL-4) in response to certain pathogens and can influence T-cell function during the development of an immune response.10 By releasing proinflammatory cytokines, T cells can also facilitate activation of other host cells such as macrophages11,12 and NK cells12-14 in the early phases of infection.
Several recent studies have focused on the role played by donor T cells in GVHD,15-21 but the role of host T cells is not known. Using several mouse models, we have found that host T cells exacerbate acute GVHD by enhancing the allostimulatory capacity of host APCs.
Materials and methods
Mice
Female C57BL/6 (B6, H-2b), BALB/c (H-2d), B6.C-H2bm12 (bm12), B6.C-H2bm1 (bm1), B6.129S7-Ifngtm1Ts/J (IFN--/-; IFN- deficient), B6.129P2-Tcrbtm1Mom/J (-/-; T-cell deficient), and B6.129P2-Tcrdtm1Mom/J (-/-; T-cell deficient) mice22,23 (both TCR-deficient mice were backcrossed onto C57BL/6 genetic background) were purchased from The Jackson Laboratory (Bar Harbor, Me). B6.SJL-Ptprca/BoAiTac-Abbtm1N13 (II-/-, B6.Ly-5a-background, major histocompatibility antigen (MHC) class II deficient) mice were purchased from Taconic Farms (Germantown, NY). B6.129P2-Tnfrsf5tm1Kik/J (CD40-/-, CD40 deficient) mice were kindly provided by Dr D. Keith Bishop. The age of mice used for experiments ranged between 8 and 12 weeks.
Bone marrow transplantation
Mice received 1100 cGy total body irradiation (TBI; 137Cs source). Bone marrow cells (5 x 106) plus 2 x 106 nylon wool-purified (NWP) splenic CD3+ T cells from respective allogeneic or syngeneic donors were resuspended in 0.25 mL Leibovitz L-15 media (Gibco BRL, Grand Island, NY) and injected intravenously into recipients on day 0. Bone marrow cells plus 1 x 106 CD4+ T cells or 2 x 106 CD8+ T cells isolated by immunomagnetic selection (Miltenyi Biotec, Bergisch Gladbach, Germany) were injected into recipients in MHC class II or I mismatched mouse model, respectively.
Clinical and histopathologic analysis of GVHD
Survival was monitored daily, and GVHD clinical scores were assessed weekly by a scoring system incorporating 5 clinical parameters: weight loss, posture (hunching), activity, fur texture, and skin integrity, as described.24 Individual mice were ear-tagged and graded weekly on a scale from 0 to 2 for each criterion (maximum score 10).
Formalin-preserved small (ileum) intestines were embedded in paraffin, cut into 5-μm thick sections, and stained with hematoxylin and eosin for histologic examination. Slides were coded without reference to prior treatment and examined in a blinded fashion by a pathologist (C.L.). A semiquantitative scoring system was used to assess the following abnormalities known to be associated with GVHD:25 villous blunting, crypt regeneration, loss of enterocyte brush border, luminal sloughing of cellular debri, crypt cell apoptosis, crypt destruction, and lamina propria lymphocytic infiltrate. The scoring system denoted 0 as normal, 0.5 as focal and rare, 1.0 as focal and mild, 2.0 as diffuse and mild, 3.0 as diffuse and moderate, and 4.0 as diffuse and severe. Scores were added to provide a total score for each specimen. After scoring the codes were broken and data compiled.
Cytokine eLISA
Concentrations of IFN-, IL-2 (BD Pharmingen, San Diego, CA), and TNF- (Quantikine; R&D Systems, Minneapolis, MN) were measured by sandwich enzyme-linked immunosorbent assay (eLISA) as described.26 To determine serum levels of lipopolysaccharide (LPS), the limulus amebocyte lysate assay (BioWhittaker, Walkersville, MD) was performed according to the manufacturer's protocol. All units expressed are relative to US reference standard eC-6.
In vivo depletion of T cells
Five days before BMT, mice were depleted of T cells as previously described27 by intraperitoneal injection of 500 μg hamster anti-TCR monoclonal antibody (mAb) (IgG, UC7-13D5; LigoCyte Pharmaceutical, Bozeman, MT) in a volume of 0.25 mL Leibovitz L-15 media. Sham depletion was accomplished with hamster Ig (Jackson Laboratories, Bar Harbor, Me).
Adoptive transfer of T-cell experiments
T-cell reconstitution was performed as described previously.28 T cells were obtained from -/- donor mice and T cells were obtained from -/- donors. Briefly, splenocytes from mice were isolated, and red blood cells were lysed. Cells were resuspended in Leibovitz L-15 media. Cells (60 x 106) were injected in the tail vein of each mouse. At 24 hours after transfer, mice received 1100 cGy TBI following BMT for clinical assessment of GVHD.
Monoclonal antibodies and blocking reagents
The following mAbs were used for flow cytometry and blocking experiments: CD3-fluorescein isothiocyanate (FITC), H2Dd-FITC, CD4-FITC, CD45.1-phycoerythrin (Pe), I-Ab-Pe, CD40-Pe, CD86-Pe, anti-CD40 ligand (L)-e, anti-TNF--Pe, anti-IFN--Pe, TCR -Pe, CD11c-allophycocyanin, CD4-allophycocyanin, CD8-allophycocyanin (all from BD PharMingen, San Diego, CA).
Cell surface phenotype and intracellular cytokine analysis
Fluorescence-activated cell sorting (FACS) analysis was performed as described previously.29 Briefly, cells were incubated with mAb 2.4G2 (rat anti-mouse FcR mAb) for 15 minutes at 4°C and then with the relevant mAb for 30 minutes at 4°C. Three-color flow cytometry was performed with an ePICS elite eSP cell sorter (Beckman Coulter, Miami, FL). For intracellular cytokine staining, splenocytes were harvested 7 days after transplantation. Cells were incubated for 4 hours with phorbol 12-myristate 13-acetate (PMA)-ionomycin (BD Pharmingen) and brefeldin A at 37°C. Then, the cells were permeabilized with the Cytofix/Cytoperm Kit from BD Pharmingen and subsequently stained with Pe-conjugated IFN- mAb. Flow cytometry was conducted by gating for the designated cell populations.
Preparation of DCs and T cells
Splenic dendritic cells (DCs) were isolated as described previously.3 Briefly, after digestion with collagenase D (1 mg/mL; Boehringer Mannheim, Indianapolis, IN), cells were resuspended in 1.035 g/mL Percoll (Pharmacia Biotech, Uppsala, Sweden) and underlaid with an equivalent volume of 1.075 g/mL Percoll. After centrifugation, the resulting band was harvested and washed twice, and DCs were isolated by using CD11c (N418) microbeads and the AutoMACS (Miltenyi Biotec).
Bone marrow-derived DCs were generated as described previously.30 Briefly, bone marrow cells were flushed from tibias and femurs of B6 mice and seeded at 2 x 106 cells in bacteriologic Petri dishes (100-mm diameter; Falcon, NJ) in a volume of 10 mL of complete medium (CM) containing 20 ng/mL granulocyte macrophage-colony-stimulating factor (GM-CSF). On day 3, 10 mL CM containing 20 ng/mL GM-CSF was added to the dishes. On day 6, nonadherent cells were harvested, washed twice, and incubated with anti-CD11c microbeads for magnetic cell separation with an AutoMACS system. The purity of the DC suspension was greater than 90% as determined by dual positivity for MHC class II (I-Ad) and CD11c.
T cells were collected from spleen in -/- mice (as in "Bone marrow transplantation") or intestinal intraepithelial lymphocytes (iIeLs) in wild-type B6 mice. iIeLs were separated as described.31 Briefly, Peyer patches were pinched out of the small intestine, which was opened longitudinally and cut into 5-mm pieces. The pieces were washed in 4°C CMF/HePeS (calcium/magnesium free N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid) and transferred into a 50-mL erlenmeyer flask with 37°C CMF/HePeS/eDTA (ethylenediaminetetraacetic acid) solution on a magnetic stirrer. After stirring, the supernatant containing iIeLs was collected and the iIeLs were enriched with a Percoll procedure. Lymphocytes were stained with Pe-labeled anti-TCR mAb. + T cells, and -depleted lymphocyte subset ( T cells) was purified by cell sorting, using a FACSVantage (BD Biosciences, San Jose, CA). Purity of + T cells assessed using flow cytometry was greater than 99%. Contamination of T cells in -depleted lymphocytes subset was less than 0.1%.
T cell-DC cocultures and mixed leukocyte reactions
T cell-DC cocultures and mixed leukocyte reactions were performed as described previously.32 Briefly, bone marrow-derived DCs were cultured in 24-well tissue-culture plates at a concentration of 1 x 106 cells/mL for 36 hours in RPMI complete medium. Additions to certain wells included 10 ng/mL escherichia coli LPS (Sigma-Aldrich, Poole, United Kingdom), + T cells (T-cell to DC ratio = 1:1), or - T cells. Anti-IFN- (10 μg/mL), anti-TNF- (20 μg/mL), and anti-CD40L (10 μg/mL) mAbs were used for blocking experiments. In some experiments, DCs were isolated after 36 hours by centrifugation on a 14.5% metrizamide (Sigma Chemical, St Louis, MO) gradient,33 irradiated (2000 cGy), and then used as stimulators for allogeneic NWP responder T cells (T-cell to DC ratio = 10:1) for 3 to 4 days. Splenic DCs isolated from mice were irradiated (1000 cGy) and then used as stimulators. At 48 and 72 hours, supernatants were collected for cytokine analysis. During the final 12 hours of culture, cells were pulsed with 1 μCi [3H] thymidine (2 Ci/mmol; New england Nuclear, Shelton, CT) and proliferation was determined on a Top Count NTX (Packard Instruments, Meriden, CT).
Statistical analysis
The Mann-Whitney U test was used for the statistical analysis of cytokine data, LPS levels, clinical scores, weight loss, and histology, whereas the Wilcoxon rank test was used to analyze survival data. P less than .05 was considered statistically significant.
Results
Absence of host T cells reduces GVHD severity
The role of host T cells on the severity of GVHD was evaluated by using a well-characterized experimental model of GVHD, BALB/c (H2d) B6 (H2b). Wild-type B6 and -/- mice were irradiated with 1100 cGy TBI and underwent transplantation with 5 x 106 BM cells and 2 x 106 splenic CD3+ T cells from either syngeneic B6 or allogeneic BALB/c donors. -/- mice that received allogeneic cells demonstrated significantly better survival than the wild-type allogeneic recipients (P < .001, Figure 1A). Clinical GVHD severity was assessed by a standard scoring system that sums changes in 5 parameters: weight loss, posture (hunching), activity, fur texture, and skin integrity (see "Materials and methods"). Consistent with improved survival, allogeneic B6 -/- BM transplant recipients demonstrated significantly less clinical acute GVHD than wt controls (P < .05, Figure 1B). All surviving transplant recipient mice in both groups displayed complete donor hematopoietic chimerism, ruling out mixed chimerism as a cause for reduced GVHD (data not shown).
We next analyzed histopathologic changes of the small intestine using a semiquantitative index for GVHD damage (see "Materials and methods"). As shown in Figure 3A-D, allogeneic B6 -/- BM transplant recipients demonstrated significantly less GVHD in gastrointestinal tract compared with wt controls (P < .05). Serum levels of LPS and TNF-, biochemical markers that correlate with the severity of intestinal and systemic acute GVHD,25,34 were also significantly reduced (Figure 3e-F). Thus -/- mice showed less acute GVHD compared with wt controls as determined by clinical, histopathologic, and biochemical markers.
To rule out strain-dependent artifacts, we next evaluated the role of host T cells in well-defined GVHD models in which donor and recipient differed at MHC class II or class I only (bm12 B6 or bm1 B6, respectively). In the CD4+-mediated, MHC class II-mismatched GVHD model (bm12 B6), -/- recipients demonstrated significantly better survival at day 35 (73% vs 16%, P < .001) and reduced clinical severity of acute GVHD on day 7 compared with wt mice (3.8 ± 0.2 vs 5.0 ± 0.1, P < .001). In the CD8+-mediated, MHC class I-mismatched model (bm1 B6), clinical GVHD was also reduced (3.1 ± 0.1 vs 4.8 ± 0.2, P < .001), although there was no significant difference in survival rate at day 50 (94% vs 73%, P = .19). Taken together, these data demonstrate that host T cells regulate CD4+- and CD8+-mediated acute GVHD.
Adoptive transfer of host T cells aggravate GVHD
To evaluate whether T cells were unique lymphocyte subsets in their ability to amplify GVHD, we compared the severity of GVHD among recipients that were deficient in either T cells or T cells. As shown in Figure 4A-B, B6 -/- recipients of allogeneic BMT showed significantly reduced GVHD mortality and clinical scores (P < .01), whereas mortality in B6 -/- recipients was similar to wt controls.
To confirm the specificity of T cell-mediated in vivo effects on GVHD severity, we reconstituted -/- mice with either T cells or T cells that were syngeneic to the host, as described in "Materials and methods." Because NK cells could activate DC function, we determined the number of NK cells in the donor inocula. Contamination of NK cells in splenocytes from both -/- and -/- donors was similar (6.5% + 0.7% vs 5.6% + 0.7%, respectively). Mice reconstituted with T cells showed significantly higher mortality and clinical scores than unreconstituted mice or mice reconstituted with T cells (Figure 4C-D). Taken together, these results demonstrate the unique ability of the host T-cell subset in the host to amplify the severity of GVHD.
T cells induce activation and allostimulatory capacity of DCs
The activation of donor T cells by host DCs is critical to the induction of GVHD.1-3 We first determined the amplitude of donor T-cell expansion and intracellular cytokine expression in the spleens of BM transplant recipients 7 days later in the BALB/c B6 model. We observed a significant (P < .01) reduction in the numbers of donor CD4+ and CD8+ T cells in -/- recipients compared with allogeneic controls as shown in Figure 5A-B. Consistent with this reduced expansion, the number of CD4+ and CD8+ donor T cells that produced IFN- was also significantly reduced (IFN-+ CD4+: 1.8 ± 0.5 vs 18.9 ± 5.0 x 105, P < .01; IFN-+ CD8+: 0.68 ± 0.15 vs 14.6 ± 4.5 x 105, P < .01).
We hypothesized that the reduced donor T-cell expansion in -/- recipients was due to a reduced allostimulatory capacity of host DCs and we thus evaluated the ability of host DCs to stimulate allogeneic responder T cells from wt and -/- mice. T cells proliferated less when cocultured with allogeneic DCs from -/- spleens than with DCs from wt spleens 6 hours after irradiation (Figure 5C, P < .05). They also secreted less IL-2 and IFN- (Figure 5D-e, P < .05).
We next analyzed the ability of T cells to enhance the stimulatory function of DCs in vitro. Intestinal intraepithelial lymphocytes (iIeLs) were isolated from B6 mice, stained with anti-TCR mAb, and sorted, producing 2 populations: + T cells (> 99% purity + by FACS) and - T cells (< 0.1% + by FACS). Bone marrow-derived DCs from B6 mice were cultured for 36 hours with either LPS, + T cells, or - T cells at a T-cell to DC ratio of 1:1. After culture, DCs were isolated by centrifugation on a 14.5% metrizamide gradient, irradiated, and then used as stimulators for allogeneic BALB/c responder T cells. DCs cultured in the presence of + T cells stimulated significantly greater proliferation (Figure 6A, P < .01) of IL-2 and IFN- secretion of allogeneic T cells when compared with DCs that had been cultured in - T cells (IL-2; 213 ± 19 pg/mL vs 62 ± 15 pg/mL, P < .01, IFN-; 5059 ± 167 pg/mL vs 1350 ± 98 pg/mL, P < .01).
We next evaluated effect of T-cell coculture on the phenotype of DCs. T cells increased the expression of CD40 from 53% to 84% (Figure 6B), whereas -depleted lymphocytes had minimal effects on CD40 (53% vs 64%). expression of MHC class II and CD80 on DCs was greater than 90% in all cultures and did not change when cocultured with T cells (data not shown).
T cells activate DCs and enhance allogeneic responses in vivo
We directly evaluated whether T cells activate DCs and enhance the responses of allogeneic T cells in vivo. T cells were depleted from B6.Ly-5a (CD45.1+) MHC class II-deficient mice (II-/-) by intraperitoneal injection of anti-TCR mAb for 5 days prior to BMT.27 These animals then received 1100 cGy TBI and were injected with 5 x 106 wt B6 DCs that had previously been cocultured either with T cells or with medium. The following day mice received 5 x 106 NWP T cells from bm12 (CD45.1-) donors. Spleens were harvested on day +7 after BMT and analyzed by FACS. The phenotype of donor CD4+ T cells was determined to be CD4+ CD45.1-. Recipients of DCs cocultured with T cells showed a 4-fold increase in donor CD4+ T cells compared with recipients of control DCs (Figure 7A). Serum levels of TNF-, a biochemical marker of GVHD, were also twice as high in these recipients (P < .05, Figure 7B). We conclude that T cells activate host DCs both in vitro and in vivo and amplify the response of allogeneic donor T cells that cause GVHD.
Cell-to-cell contact is required for T-cell-mediated DC activation
Because T cells can secrete activating cytokines such as IFN-,10,12,13,35,36 we examined a potential role for cytokines in the enhancement of the allostimulatory capacity of DCs caused by these cells. As shown in Figure 6C, the addition of anti-IFN- mAb to coculture did not inhibit the activation of DCs by T cells. DCs matured in the presence of T cells from IFN--/- mice stimulated allogeneic T cells to the same degree as DCs from wt mice (Figure 6C). Similar results were observed with anti-TNF- mAb (data not shown). The percentage of DCs expressing CD40 was increased when cocultured with T cells. CD40L is a costimulatory molecule that is upregulated on CD4+ T cells after exposure to antigen and plays a critical role in DC activation.37-40 However, mAb blockade of CD40L did not prevent DC activation (Figure 6e). Furthermore, DCs from CD40-/- mice were activated to the same degree as DCs from wt mice, confirming that CD40-CD40L interactions are not required for T-cell-mediated DC activation (Figure 6D).
We next determined whether cell-to-cell contact was necessary for the T-cell activation of DCs. DCs were cultured for 36 hours either in direct contact with T cells or separated by a 0.2 μ transwell membrane and were then used as stimulators of allogeneic T cells. Physical separation of DCs and T cells in the transwells completely abrogated DC activation (Figure 6e), demonstrating the requirement for cell-to-cell contact.
Discussion
The localization of T cells within GVHD target tissues and their ability to modulate immune responses suggested that these cells might play a role in the pathogenesis of GVHD. In this study we demonstrate that the absence of T cells in allogeneic BM transplant recipients caused significantly less severe acute GVHD as determined by all measurable parameters in several different donor/recipient strain combinations. This finding was also confirmed by depleting T cells in wt recipients by in vivo administration of mAb against TCRs. Reconstitution of host T cells to -/- recipients increased lethal GVHD. Coculture of T cells with DCs enhanced their allostimulatory capacity and increased the proliferation of allogeneic responder T cells in vitro and in vivo. This enhancement was independent of IFN-, TNF-, and CD40L, but depended on cell-to-cell contact. These data demonstrate that host T cells play a critical role in the pathogenesis of GVHD by amplifying the stimulatory function of host DCs.
Donor T cells' interactions with host DCs are essential for the induction of GVHD.1-3 DCs represent a pool of professional APCs whose maturation/activation state is critical to their function. The chemo-radiotherapy used to condition BM transplant recipients induces the secretion of inflammatory cytokine such as IL-1, TNF-, and IL-6 from damaged host tissues,41 and allows LPS to enter into the systemic circulation.42 Such "danger signals" are critical for the activation of host DCs,43 but Shlomchik44 has reported that recipients deficient in TNFR1/R2, IL-1R, CD40, and Toll-like receptor 4 (essential for LPS-induced activation of DCs) developed GVHD comparable to that seen in wild-type controls. These experiments exclude a role for each of these individual pathways in the activation of host DCs and suggest that either together or through other pathways and/or cell interactions might contribute to the process. Several cell types such as macrophages,5 NK cells,45 CD4+ T,37-40 and CD8+ T46 have been shown to provide maturation and activation signals for DCs.
Our data demonstrate that murine T cells promote DC activation in a cell-to-cell contact-dependent manner. Leslie et al32 have also demonstrated that human T cells promote DC maturation in a cell-to-cell contact-dependent manner that is restricted by CD1c. Mice do not express CD1c but do express only CD1d, which presents glycolipid antigens to NKT cells.47 Recently, Hashimoto et al48 have demonstrated that absence of CD1d in BM transplant recipients results in a loss of protection from GVHD that is mediated by NKT cells. Thus, while our data demonstrated that T cells exacerbate GVHD, NKT cells can reduce GVHD, suggesting that in mice CD1 probably regulates GVHD through its interaction with NKT cells.
Other studies have focused on the role of T cells in the pathogenesis of GVHD.15-21 One study reported that a clonal population of T cells can induce lethal GVHD,17 whereas Drobyski and Majewski20 and Drobyski et al21 demonstrated that donor T cells alone do not cause acute GVHD. Coadministration of T cells and T cells exacerbated GVHD when compared with T cells alone, suggesting that donor T cells could contribute to GVHD but required the presence of T cells.18 However, the role played by host T cells in the pathogenesis of allogeneic GVHD has not been defined. Sakai et al27 reported that allogeneic BMT recipients depleted of T cells by in vivo administration of mAb against TCRs showed less intestinal damage, but the mechanisms for this phenomenon were not explored. We demonstrate that host T cells amplify systemic GVHD in irradiated GVHD models. Our data suggest that they might function as the "danger signals" and activate the DCs, thus enhancing their allostimulatory capacity.
Although T cells are a minor subpopulation of lymphocytes within lymph nodes and the spleen, they predominate in the epithelium of several principal target organs of GVHD, such as the skin, intestinal tract, and lung.49 T cells respond to infected or transformed epithelial cells and may therefore function as sensors of danger signals that can activate an immune response.43 Recent evidence suggests that IeLs express several chemokines such as macrophage inflammatory protein 1 (MIP1), MIP1, Regulated or Activation, Normal-T cell expressed and Secreted (RANTeS), and lymphotactin, and therefore play an important role in the rapid recruitment of CCR5+ cells into these epithelial tissues, helping to explain the target organ distribution of GVHD.50,51
T cells may also affect events in the gastrointestinal (GI) tract through their production of keratinocyte growth factor (KGF).52 KGF administration can reduce the severity of GVHD, particularly in the GI tract, and GVHD mortality.53,54 An absence of T cells would be expected to result in reduced endogenous KGF production and enhanced GI tract damage. To examine this possibility we compared the degree of tissue damage on day 7 after lethal irradiation and found no significant difference between wild-type and -/- mice (data not shown). Mice devoid of T cells showed decreased histopathologic damage from GVHD after allogeneic BMT (Figure 3), suggesting that whatever potential benefit T cells might have in GVHD by the secretion of KGF and subsequent promotion of tissue repair, it is overshadowed by the activation of host APCs and a greater donor response of allogeneic T cells.
In summary, we demonstrate a novel role for host T cells in the amplification of GVHD through the activation of host APCs, suggesting that elimination of host T cells might be a novel therapeutic strategy to reduce GVHD.
Footnotes
Prepublished online as Blood First edition Paper, March 29, 2005; DOI 10.1182/blood-2004-10-4087.
Supported by National Institutes of Health grants K08 AI052 863-01 (P.R.), DK 02 958 (C.L.), and CA 49 542 (J.L.M.F.). J.L.M.F. is a Doris Duke Distinguished Clinical Scientist.
An Inside Blood analysis of this article appears in the front of this issue.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
References
Zhang Y, Louboutin JP, Zhu J, Rivera AJ, emerson SG. Preterminal host dendritic cells in irradiated mice prime CD8+ T cell-mediated acute graft-versus-host disease. J Clin Invest. 2002;109: 1335-1344.
Shlomchik WD, Couzens MS, Tang CB, et al. Prevention of graft versus host disease by inactivation of host antigen-presenting cells. Science. 1999;285: 412-415.
Duffner UA, Maeda Y, Cooke KR, et al. Host dendritic cells alone are sufficient to initiate acute graft-versus-host disease. J Immunol. 2004;172: 7393-7398.
Mowat AM, Viney JL. The anatomical basis of intestinal immunity. Immunol Rev. 1997;156: 145-166.
Nestel FP, Price KS, Seemayer TA, Lapp WS. Macrophage priming and lipopolysaccharide-triggered release of tumor necrosis factor alpha during graft-versus-host disease. J exp Med. 1992;175: 405-413.
Ruggeri L, Capanni M, Urbani e, et al. effectiveness of donor natural killer cell alloreactivity in mismatched hematopoietic transplants. Science. 2002;295: 2097-2100.
Bryson JS, Flanagan DL. Role of natural killer cells in the development of graft-versus-host disease. J Hematother Stem Cell Res. 2000;9: 307-316.
Zeng D, Lewis D, Dejbakhsh-Jones S, et al. Bone marrow NK1.1(-) and NK1.1(+) T cells reciprocally regulate acute graft versus host disease. J exp Med. 1999;189: 1073-1081.
Chien YH, Hampl J. Antigen-recognition properties of murine gamma delta T cells. Springer Semin Immunopathol. 2000;22: 239-250.
Ferrick DA, Schrenzel MD, Mulvania T, Hsieh B, Ferlin WG, Lepper H. Differential production of interferon-gamma and interleukin-4 in response to Th1- and Th2-stimulating pathogens by gamma delta T cells in vivo. Nature. 1995;373: 255-257.
Jones-Carson J, Vazquez-Torres A, van der Heyde HC, Warner T, Wagner RD, Balish e. Gamma delta T cell-induced nitric oxide production enhances resistance to mucosal candidiasis. Nat Med. 1995;1: 552-557.
Skeen MJ, Ziegler HK. Activation of gamma delta T cells for production of IFN-gamma is mediated by bacteria via macrophage-derived cytokines IL-1 and IL-12. J Immunol. 1995;154: 5832-5841.
Nishimura H, emoto M, Hiromatsu K, et al. The role of gamma delta T cells in priming macrophages to produce tumor necrosis factor-alpha. eur J Immunol. 1995;25: 1465-1468.
Ladel CH, Blum C, Kaufmann SH. Control of natural killer cell-mediated innate resistance against the intracellular pathogen Listeria monocytogenes by gamma/delta T lymphocytes. Infect Immun. 1996;64: 1744-1749.
ellison CA, MacDonald GC, Rector eS, Gartner JG. Gamma delta T cells in the pathobiology of murine acute graft-versus-host disease: evidence that gamma delta T cells mediate natural killer-like cytotoxicity in the host and that elimination of these cells from donors significantly reduces mortality. J Immunol. 1995;155: 4189-4198.
Huang Y, Cramer De, Ray MB, Chilton PM, Que X, Ildstad ST. The role of alphabeta- and gammadelta-T cells in allogenic donor marrow on engraftment, chimerism, and graft-versus-host disease. Transplantation. 2001;72: 1907-1914.
Blazar BR, Taylor PA, Panoskaltsis-Mortari A, Barrett TA, Bluestone JA, Vallera DA. Lethal murine graft-versus-host disease induced by donor gamma/delta expressing T cells with specificity for host nonclassical major histocompatibility complex class Ib antigens. Blood. 1996;87: 827-837.
Drobyski WR, Vodanovic-Jankovic S, Klein J. Adoptively transferred gamma delta T cells indirectly regulate murine graft-versus-host reactivity following donor leukocyte infusion therapy in mice. J Immunol. 2000;165: 1634-1640.
Tsuji S, Char D, Bucy RP, Simonsen M, Chen CH, Cooper MD. Gamma delta T cells are secondary participants in acute graft-versus-host reactions initiated by CD4+ alpha beta T cells. eur J Immunol. 1996;26: 420-427.
Drobyski WR, Majewski D. Donor gamma delta T lymphocytes promote allogeneic engraftment across the major histocompatibility barrier in mice. Blood. 1997;89: 1100-1109.
Drobyski WR, Majewski D, Hanson G. Graft-facilitating doses of ex vivo activated gammadelta T cells do not cause lethal murine graft-vs-host disease. Biol Blood Marrow Transplant. 1999;5: 222-230.
Mombaerts P, Clarke AR, Rudnicki MA, et al. Mutations in T-cell antigen receptor genes alpha and beta block thymocyte development at different stages. Nature. 1992;360: 225-231.
Mombaerts P, Arnoldi J, Russ F, Tonegawa S, Kaufmann SH. Different roles of alpha beta and gamma delta T cells in immunity against an intracellular bacterial pathogen. Nature. 1993;365: 53-56.
Cooke KR, Kobzik L, Martin TR, et al. An experimental model of idiopathic pneumonia syndrome after bone marrow transplantation, I: the roles of minor H antigens and endotoxin. Blood. 1996;88: 3230-3239.
Cooke KR, Hill GR, Crawford JM, et al. Tumor necrosis factor-alpha production to lipopolysaccharide stimulation by donor cells predicts the severity of experimental acute graft-versus-host disease. J Clin Invest. 1998;102: 1882-1891.
Teshima T, Mach N, Hill GR, et al. Tumor cell vaccine elicits potent antitumor immunity after allogeneic T-cell-depleted bone marrow transplantation. Cancer Res. 2001;61: 162-171.
Sakai T, Ohara-Inagaki K, Tsuzuki T, Yoshikai Y. Host intestinal intraepithelial gamma delta T lymphocytes present during acute graft-versus-host disease in mice may contribute to the development of enteropathy. eur J Immunol. 1995;25: 87-91.
Wang T, Scully e, Yin Z, et al. IFN-gamma-producing gamma delta T cells help control murine West Nile virus infection. J Immunol. 2003;171: 2524-2531.
Hill GR, Crawford JM, Cooke KR, Brinson YS, Pan L, Ferrara JL. Total body irradiation and acute graft-versus-host disease: the role of gastrointestinal damage and inflammatory cytokines. Blood. 1997;90: 3204-3213.
Lutz MB, Kukutsch N, Ogilvie AL, et al. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J Immunol Methods. 1999;223: 77-92.
Duffner U, Lu B, Hildebrandt GC, et al. Role of CXCR3-induced donor T-cell migration in acute GVHD. exp Hematol. 2003;31: 897-902.
Leslie DS, Vincent MS, Spada FM, et al. CD1-mediated gamma/delta T cell maturation of dendritic cells. J exp Med. 2002;196: 1575-1584.
Fields RC, Shimizu K, Mule JJ. Murine dendritic cells pulsed with whole tumor lysates mediate potent antitumor immune responses in vitro and in vivo. Proc Natl Acad Sci U S A. 1998;95: 9482-9487.
Hill GR, Ferrara JL. The primacy of the gastrointestinal tract as a target organ of acute graft-versus-host disease: rationale for the use of cytokine shields in allogeneic bone marrow transplantation. Blood. 2000;95: 2754-2759.
Moore TA, Moore BB, Newstead MW, Standiford TJ. Gamma delta-T cells are critical for survival and early proinflammatory cytokine gene expression during murine Klebsiella pneumonia. J Immunol. 2000;165: 2643-2650.
Gao Y, Yang W, Pan M, et al. Gamma delta T cells provide an early source of interferon gamma in tumor immunity. J exp Med. 2003;198: 433-442.
Ridge JP, Di Rosa F, Matzinger P. A conditioned dendritic cell can be a temporal bridge between a CD4+ T-helper and a T-killer cell. Nature. 1998;393: 474-478.
Bennett SR, Carbone FR, Karamalis F, Flavell RA, Miller JF, Heath WR. Help for cytotoxic-T-cell responses is mediated by CD40 signalling. Nature. 1998;393: 478-480.
Schoenberger SP, Toes Re, van der Voort eI, Offringa R, Melief CJ. T-cell help for cytotoxic T lymphocytes is mediated by CD40-CD40L interactions. Nature. 1998;393: 480-483.
Cella M, Scheidegger D, Palmer-Lehmann K, Lane P, Lanzavecchia A, Alber G. Ligation of CD40 on dendritic cells triggers production of high levels of interleukin-12 and enhances T cell stimulatory capacity: T-T help via APC activation. J exp Med. 1996;184: 747-752.
Xun CQ, Thompson JS, Jennings CD, Brown SA, Widmer MB. effect of total body irradiation, busul-fan-cyclophosphamide, or cyclophosphamide conditioning on inflammatory cytokine release and development of acute and chronic graft-versus-host disease in H-2-incompatible transplanted SCID mice. Blood. 1994;83: 2360-2367.
Ferrara JL, Cooke KR, Teshima T. The pathophysiology of acute graft-versus-host disease. Int J Hematol. 2003;78: 181-187.
Gallucci S, Matzinger P. Danger signals: SOS to the immune system. Curr Opin Immunol. 2001;13: 114-119.
Shlomchik WD. Antigen presentation in graft-vs-host disease. exp Hematol. 2003;31: 1187-1197.
Zitvogel L. Dendritic and natural killer cells cooperate in the control/switch of innate immunity. J exp Med. 2002;195: F9-14.
Ruedl C, Kopf M, Bachmann MF. CD8(+) T cells mediate CD40-independent maturation of dendritic cells in vivo. J exp Med. 1999;189: 1875-1884.
Gumperz Je, Brenner MB. CD1-specific T cells in microbial immunity. Curr Opin Immunol. 2001;13: 471-478.
Hashimoto D, Asakura S, Miyake S, et al. Host NKT cells promote Th2 polarization of donor T cells and regulate acute GVHD after experimental BMT via a STAT6-dependent mechanism. Blood. 2003;102: 191A-192A.
Augustin A, Kubo RT, Sim GK. Resident pulmonary lymphocytes expressing the gamma/delta T-cell receptor. Nature. 1989;340: 239-241.
Shires J, Theodoridis e, Hayday AC. Biological insights into TCRgammadelta+ and TCRalpha-beta+ intraepithelial lymphocytes provided by serial analysis of gene expression (SAGe). Immunity. 2001;15: 419-434.
Boismenu R, Feng L, Xia YY, Chang JC, Havran WL. Chemokine expression by intraepithelial gamma delta T cells: implications for the recruitment of inflammatory cells to damaged epithelia. J Immunol. 1996;157: 985-992.
Boismenu R, Havran WL. Modulation of epithelial cell growth by intraepithelial gamma delta T cells. Science. 1994;266: 1253-1255.
Panoskaltsis-Mortari A, Lacey DL, Vallera DA, Blazar BR. Keratinocyte growth factor administered before conditioning ameliorates graft-versus-host disease after allogeneic bone marrow transplantation in mice. Blood. 1998;92: 3960-3967.
Krijanovski OI, Hill GR, Cooke KR, et al. Keratinocyte growth factor separates graft-versus-leukemia effects from graft-versus-host disease. Blood. 1999;94: 825-831.(Yoshinobu Maeda, Pavan Re)
Pediatrics, University of Michigan Cancer Center, Ann Arbor, MI
the Department of Pathology, University of Florida College of Medicine, Gainesville.
Abstract
T cells localize to target tissues of graft-versus-host disease (GVHD) and therefore we investigated the role of host T cells in the pathogenesis of acute GVHD in several well-characterized allogeneic bone marrow transplantation (BMT) models. Depletion of host T cells in wild-type (wt) B6 recipients by administration of anti-T-cell receptor (TCR) monoclonal antibody reduced GVHD, and T-cell-deficient (-/-) BM transplant recipients experienced markedly improved survival compared with normal controls (63% vs 10%, P < .001). T cells were responsible for this difference because reconstitution of -/- recipients with T cells restored GVHD mortality. -/- recipients showed decreased serum levels of tumor necrosis factor (TNF-), less GVHD histopathologic damage, and reduced donor T-cell expansion. Mechanistic analysis of this phenomenon demonstrated that dendritic cells (DCs) from -/- recipients exhibited less allostimulatory capacity compared to wt DCs after irradiation. Normal DCs derived from BM caused greater allogeneic T-cell proliferation when cocultured with T cells than DCs cocultured with medium alone. This enhancement did not depend on interferon (IFN-), TNF-, or CD40 ligand but did depend on cell-to-cell contact. These data demonstrated that the host T cells exacerbate GVHD by enhancing the allostimulatory capacity of host antigen-presenting cells. (Blood. 2005;106:749-755)
Introduction
Allogeneic bone marrow transplantation (BMT) is a widely performed therapy for many hematologic malignancies. However, acute graft-versus-host disease (GVHD) remains the most important complication of allogeneic BMT. The pathophysiology of GVHD is complex. The target organs of acute GVHD include the gastrointestinal tract, skin, and liver. Several convergent lines of experimental data have demonstrated that donor T cells and host antigen-presenting cells (APCs) are critical for the induction of GVHD.1-3 In addition, a growing body of data also suggest that several other cellular subsets such as host macrophages,4,5 donor natural killer (NK) cells,6,7 and donor NKT cells8 are involved in the pathogenesis of GVHD.
T cells represent a distinct lineage of T cells that evolved early in vertebrate phylogeny in conjunction with T cells and B cells. Antigen recognition by T-cell receptors (TCRs) occurs in a fundamentally different manner from that of T cells. T cells can recognize molecules other than peptides, do not require professional APCs for activation, and use their TCRs to recognize patterns associated with foreign pathogens or tissue injury.9 These cells serve as sentinels to monitor their environment for signs of stress caused by infection, inflammation, and transformation. T cells modulate immune responses by secreting cytokines and by regulating the function of other immune cells. For example, T cells produce either interferon (IFN-) or interleukin 4 (IL-4) in response to certain pathogens and can influence T-cell function during the development of an immune response.10 By releasing proinflammatory cytokines, T cells can also facilitate activation of other host cells such as macrophages11,12 and NK cells12-14 in the early phases of infection.
Several recent studies have focused on the role played by donor T cells in GVHD,15-21 but the role of host T cells is not known. Using several mouse models, we have found that host T cells exacerbate acute GVHD by enhancing the allostimulatory capacity of host APCs.
Materials and methods
Mice
Female C57BL/6 (B6, H-2b), BALB/c (H-2d), B6.C-H2bm12 (bm12), B6.C-H2bm1 (bm1), B6.129S7-Ifngtm1Ts/J (IFN--/-; IFN- deficient), B6.129P2-Tcrbtm1Mom/J (-/-; T-cell deficient), and B6.129P2-Tcrdtm1Mom/J (-/-; T-cell deficient) mice22,23 (both TCR-deficient mice were backcrossed onto C57BL/6 genetic background) were purchased from The Jackson Laboratory (Bar Harbor, Me). B6.SJL-Ptprca/BoAiTac-Abbtm1N13 (II-/-, B6.Ly-5a-background, major histocompatibility antigen (MHC) class II deficient) mice were purchased from Taconic Farms (Germantown, NY). B6.129P2-Tnfrsf5tm1Kik/J (CD40-/-, CD40 deficient) mice were kindly provided by Dr D. Keith Bishop. The age of mice used for experiments ranged between 8 and 12 weeks.
Bone marrow transplantation
Mice received 1100 cGy total body irradiation (TBI; 137Cs source). Bone marrow cells (5 x 106) plus 2 x 106 nylon wool-purified (NWP) splenic CD3+ T cells from respective allogeneic or syngeneic donors were resuspended in 0.25 mL Leibovitz L-15 media (Gibco BRL, Grand Island, NY) and injected intravenously into recipients on day 0. Bone marrow cells plus 1 x 106 CD4+ T cells or 2 x 106 CD8+ T cells isolated by immunomagnetic selection (Miltenyi Biotec, Bergisch Gladbach, Germany) were injected into recipients in MHC class II or I mismatched mouse model, respectively.
Clinical and histopathologic analysis of GVHD
Survival was monitored daily, and GVHD clinical scores were assessed weekly by a scoring system incorporating 5 clinical parameters: weight loss, posture (hunching), activity, fur texture, and skin integrity, as described.24 Individual mice were ear-tagged and graded weekly on a scale from 0 to 2 for each criterion (maximum score 10).
Formalin-preserved small (ileum) intestines were embedded in paraffin, cut into 5-μm thick sections, and stained with hematoxylin and eosin for histologic examination. Slides were coded without reference to prior treatment and examined in a blinded fashion by a pathologist (C.L.). A semiquantitative scoring system was used to assess the following abnormalities known to be associated with GVHD:25 villous blunting, crypt regeneration, loss of enterocyte brush border, luminal sloughing of cellular debri, crypt cell apoptosis, crypt destruction, and lamina propria lymphocytic infiltrate. The scoring system denoted 0 as normal, 0.5 as focal and rare, 1.0 as focal and mild, 2.0 as diffuse and mild, 3.0 as diffuse and moderate, and 4.0 as diffuse and severe. Scores were added to provide a total score for each specimen. After scoring the codes were broken and data compiled.
Cytokine eLISA
Concentrations of IFN-, IL-2 (BD Pharmingen, San Diego, CA), and TNF- (Quantikine; R&D Systems, Minneapolis, MN) were measured by sandwich enzyme-linked immunosorbent assay (eLISA) as described.26 To determine serum levels of lipopolysaccharide (LPS), the limulus amebocyte lysate assay (BioWhittaker, Walkersville, MD) was performed according to the manufacturer's protocol. All units expressed are relative to US reference standard eC-6.
In vivo depletion of T cells
Five days before BMT, mice were depleted of T cells as previously described27 by intraperitoneal injection of 500 μg hamster anti-TCR monoclonal antibody (mAb) (IgG, UC7-13D5; LigoCyte Pharmaceutical, Bozeman, MT) in a volume of 0.25 mL Leibovitz L-15 media. Sham depletion was accomplished with hamster Ig (Jackson Laboratories, Bar Harbor, Me).
Adoptive transfer of T-cell experiments
T-cell reconstitution was performed as described previously.28 T cells were obtained from -/- donor mice and T cells were obtained from -/- donors. Briefly, splenocytes from mice were isolated, and red blood cells were lysed. Cells were resuspended in Leibovitz L-15 media. Cells (60 x 106) were injected in the tail vein of each mouse. At 24 hours after transfer, mice received 1100 cGy TBI following BMT for clinical assessment of GVHD.
Monoclonal antibodies and blocking reagents
The following mAbs were used for flow cytometry and blocking experiments: CD3-fluorescein isothiocyanate (FITC), H2Dd-FITC, CD4-FITC, CD45.1-phycoerythrin (Pe), I-Ab-Pe, CD40-Pe, CD86-Pe, anti-CD40 ligand (L)-e, anti-TNF--Pe, anti-IFN--Pe, TCR -Pe, CD11c-allophycocyanin, CD4-allophycocyanin, CD8-allophycocyanin (all from BD PharMingen, San Diego, CA).
Cell surface phenotype and intracellular cytokine analysis
Fluorescence-activated cell sorting (FACS) analysis was performed as described previously.29 Briefly, cells were incubated with mAb 2.4G2 (rat anti-mouse FcR mAb) for 15 minutes at 4°C and then with the relevant mAb for 30 minutes at 4°C. Three-color flow cytometry was performed with an ePICS elite eSP cell sorter (Beckman Coulter, Miami, FL). For intracellular cytokine staining, splenocytes were harvested 7 days after transplantation. Cells were incubated for 4 hours with phorbol 12-myristate 13-acetate (PMA)-ionomycin (BD Pharmingen) and brefeldin A at 37°C. Then, the cells were permeabilized with the Cytofix/Cytoperm Kit from BD Pharmingen and subsequently stained with Pe-conjugated IFN- mAb. Flow cytometry was conducted by gating for the designated cell populations.
Preparation of DCs and T cells
Splenic dendritic cells (DCs) were isolated as described previously.3 Briefly, after digestion with collagenase D (1 mg/mL; Boehringer Mannheim, Indianapolis, IN), cells were resuspended in 1.035 g/mL Percoll (Pharmacia Biotech, Uppsala, Sweden) and underlaid with an equivalent volume of 1.075 g/mL Percoll. After centrifugation, the resulting band was harvested and washed twice, and DCs were isolated by using CD11c (N418) microbeads and the AutoMACS (Miltenyi Biotec).
Bone marrow-derived DCs were generated as described previously.30 Briefly, bone marrow cells were flushed from tibias and femurs of B6 mice and seeded at 2 x 106 cells in bacteriologic Petri dishes (100-mm diameter; Falcon, NJ) in a volume of 10 mL of complete medium (CM) containing 20 ng/mL granulocyte macrophage-colony-stimulating factor (GM-CSF). On day 3, 10 mL CM containing 20 ng/mL GM-CSF was added to the dishes. On day 6, nonadherent cells were harvested, washed twice, and incubated with anti-CD11c microbeads for magnetic cell separation with an AutoMACS system. The purity of the DC suspension was greater than 90% as determined by dual positivity for MHC class II (I-Ad) and CD11c.
T cells were collected from spleen in -/- mice (as in "Bone marrow transplantation") or intestinal intraepithelial lymphocytes (iIeLs) in wild-type B6 mice. iIeLs were separated as described.31 Briefly, Peyer patches were pinched out of the small intestine, which was opened longitudinally and cut into 5-mm pieces. The pieces were washed in 4°C CMF/HePeS (calcium/magnesium free N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid) and transferred into a 50-mL erlenmeyer flask with 37°C CMF/HePeS/eDTA (ethylenediaminetetraacetic acid) solution on a magnetic stirrer. After stirring, the supernatant containing iIeLs was collected and the iIeLs were enriched with a Percoll procedure. Lymphocytes were stained with Pe-labeled anti-TCR mAb. + T cells, and -depleted lymphocyte subset ( T cells) was purified by cell sorting, using a FACSVantage (BD Biosciences, San Jose, CA). Purity of + T cells assessed using flow cytometry was greater than 99%. Contamination of T cells in -depleted lymphocytes subset was less than 0.1%.
T cell-DC cocultures and mixed leukocyte reactions
T cell-DC cocultures and mixed leukocyte reactions were performed as described previously.32 Briefly, bone marrow-derived DCs were cultured in 24-well tissue-culture plates at a concentration of 1 x 106 cells/mL for 36 hours in RPMI complete medium. Additions to certain wells included 10 ng/mL escherichia coli LPS (Sigma-Aldrich, Poole, United Kingdom), + T cells (T-cell to DC ratio = 1:1), or - T cells. Anti-IFN- (10 μg/mL), anti-TNF- (20 μg/mL), and anti-CD40L (10 μg/mL) mAbs were used for blocking experiments. In some experiments, DCs were isolated after 36 hours by centrifugation on a 14.5% metrizamide (Sigma Chemical, St Louis, MO) gradient,33 irradiated (2000 cGy), and then used as stimulators for allogeneic NWP responder T cells (T-cell to DC ratio = 10:1) for 3 to 4 days. Splenic DCs isolated from mice were irradiated (1000 cGy) and then used as stimulators. At 48 and 72 hours, supernatants were collected for cytokine analysis. During the final 12 hours of culture, cells were pulsed with 1 μCi [3H] thymidine (2 Ci/mmol; New england Nuclear, Shelton, CT) and proliferation was determined on a Top Count NTX (Packard Instruments, Meriden, CT).
Statistical analysis
The Mann-Whitney U test was used for the statistical analysis of cytokine data, LPS levels, clinical scores, weight loss, and histology, whereas the Wilcoxon rank test was used to analyze survival data. P less than .05 was considered statistically significant.
Results
Absence of host T cells reduces GVHD severity
The role of host T cells on the severity of GVHD was evaluated by using a well-characterized experimental model of GVHD, BALB/c (H2d) B6 (H2b). Wild-type B6 and -/- mice were irradiated with 1100 cGy TBI and underwent transplantation with 5 x 106 BM cells and 2 x 106 splenic CD3+ T cells from either syngeneic B6 or allogeneic BALB/c donors. -/- mice that received allogeneic cells demonstrated significantly better survival than the wild-type allogeneic recipients (P < .001, Figure 1A). Clinical GVHD severity was assessed by a standard scoring system that sums changes in 5 parameters: weight loss, posture (hunching), activity, fur texture, and skin integrity (see "Materials and methods"). Consistent with improved survival, allogeneic B6 -/- BM transplant recipients demonstrated significantly less clinical acute GVHD than wt controls (P < .05, Figure 1B). All surviving transplant recipient mice in both groups displayed complete donor hematopoietic chimerism, ruling out mixed chimerism as a cause for reduced GVHD (data not shown).
We next analyzed histopathologic changes of the small intestine using a semiquantitative index for GVHD damage (see "Materials and methods"). As shown in Figure 3A-D, allogeneic B6 -/- BM transplant recipients demonstrated significantly less GVHD in gastrointestinal tract compared with wt controls (P < .05). Serum levels of LPS and TNF-, biochemical markers that correlate with the severity of intestinal and systemic acute GVHD,25,34 were also significantly reduced (Figure 3e-F). Thus -/- mice showed less acute GVHD compared with wt controls as determined by clinical, histopathologic, and biochemical markers.
To rule out strain-dependent artifacts, we next evaluated the role of host T cells in well-defined GVHD models in which donor and recipient differed at MHC class II or class I only (bm12 B6 or bm1 B6, respectively). In the CD4+-mediated, MHC class II-mismatched GVHD model (bm12 B6), -/- recipients demonstrated significantly better survival at day 35 (73% vs 16%, P < .001) and reduced clinical severity of acute GVHD on day 7 compared with wt mice (3.8 ± 0.2 vs 5.0 ± 0.1, P < .001). In the CD8+-mediated, MHC class I-mismatched model (bm1 B6), clinical GVHD was also reduced (3.1 ± 0.1 vs 4.8 ± 0.2, P < .001), although there was no significant difference in survival rate at day 50 (94% vs 73%, P = .19). Taken together, these data demonstrate that host T cells regulate CD4+- and CD8+-mediated acute GVHD.
Adoptive transfer of host T cells aggravate GVHD
To evaluate whether T cells were unique lymphocyte subsets in their ability to amplify GVHD, we compared the severity of GVHD among recipients that were deficient in either T cells or T cells. As shown in Figure 4A-B, B6 -/- recipients of allogeneic BMT showed significantly reduced GVHD mortality and clinical scores (P < .01), whereas mortality in B6 -/- recipients was similar to wt controls.
To confirm the specificity of T cell-mediated in vivo effects on GVHD severity, we reconstituted -/- mice with either T cells or T cells that were syngeneic to the host, as described in "Materials and methods." Because NK cells could activate DC function, we determined the number of NK cells in the donor inocula. Contamination of NK cells in splenocytes from both -/- and -/- donors was similar (6.5% + 0.7% vs 5.6% + 0.7%, respectively). Mice reconstituted with T cells showed significantly higher mortality and clinical scores than unreconstituted mice or mice reconstituted with T cells (Figure 4C-D). Taken together, these results demonstrate the unique ability of the host T-cell subset in the host to amplify the severity of GVHD.
T cells induce activation and allostimulatory capacity of DCs
The activation of donor T cells by host DCs is critical to the induction of GVHD.1-3 We first determined the amplitude of donor T-cell expansion and intracellular cytokine expression in the spleens of BM transplant recipients 7 days later in the BALB/c B6 model. We observed a significant (P < .01) reduction in the numbers of donor CD4+ and CD8+ T cells in -/- recipients compared with allogeneic controls as shown in Figure 5A-B. Consistent with this reduced expansion, the number of CD4+ and CD8+ donor T cells that produced IFN- was also significantly reduced (IFN-+ CD4+: 1.8 ± 0.5 vs 18.9 ± 5.0 x 105, P < .01; IFN-+ CD8+: 0.68 ± 0.15 vs 14.6 ± 4.5 x 105, P < .01).
We hypothesized that the reduced donor T-cell expansion in -/- recipients was due to a reduced allostimulatory capacity of host DCs and we thus evaluated the ability of host DCs to stimulate allogeneic responder T cells from wt and -/- mice. T cells proliferated less when cocultured with allogeneic DCs from -/- spleens than with DCs from wt spleens 6 hours after irradiation (Figure 5C, P < .05). They also secreted less IL-2 and IFN- (Figure 5D-e, P < .05).
We next analyzed the ability of T cells to enhance the stimulatory function of DCs in vitro. Intestinal intraepithelial lymphocytes (iIeLs) were isolated from B6 mice, stained with anti-TCR mAb, and sorted, producing 2 populations: + T cells (> 99% purity + by FACS) and - T cells (< 0.1% + by FACS). Bone marrow-derived DCs from B6 mice were cultured for 36 hours with either LPS, + T cells, or - T cells at a T-cell to DC ratio of 1:1. After culture, DCs were isolated by centrifugation on a 14.5% metrizamide gradient, irradiated, and then used as stimulators for allogeneic BALB/c responder T cells. DCs cultured in the presence of + T cells stimulated significantly greater proliferation (Figure 6A, P < .01) of IL-2 and IFN- secretion of allogeneic T cells when compared with DCs that had been cultured in - T cells (IL-2; 213 ± 19 pg/mL vs 62 ± 15 pg/mL, P < .01, IFN-; 5059 ± 167 pg/mL vs 1350 ± 98 pg/mL, P < .01).
We next evaluated effect of T-cell coculture on the phenotype of DCs. T cells increased the expression of CD40 from 53% to 84% (Figure 6B), whereas -depleted lymphocytes had minimal effects on CD40 (53% vs 64%). expression of MHC class II and CD80 on DCs was greater than 90% in all cultures and did not change when cocultured with T cells (data not shown).
T cells activate DCs and enhance allogeneic responses in vivo
We directly evaluated whether T cells activate DCs and enhance the responses of allogeneic T cells in vivo. T cells were depleted from B6.Ly-5a (CD45.1+) MHC class II-deficient mice (II-/-) by intraperitoneal injection of anti-TCR mAb for 5 days prior to BMT.27 These animals then received 1100 cGy TBI and were injected with 5 x 106 wt B6 DCs that had previously been cocultured either with T cells or with medium. The following day mice received 5 x 106 NWP T cells from bm12 (CD45.1-) donors. Spleens were harvested on day +7 after BMT and analyzed by FACS. The phenotype of donor CD4+ T cells was determined to be CD4+ CD45.1-. Recipients of DCs cocultured with T cells showed a 4-fold increase in donor CD4+ T cells compared with recipients of control DCs (Figure 7A). Serum levels of TNF-, a biochemical marker of GVHD, were also twice as high in these recipients (P < .05, Figure 7B). We conclude that T cells activate host DCs both in vitro and in vivo and amplify the response of allogeneic donor T cells that cause GVHD.
Cell-to-cell contact is required for T-cell-mediated DC activation
Because T cells can secrete activating cytokines such as IFN-,10,12,13,35,36 we examined a potential role for cytokines in the enhancement of the allostimulatory capacity of DCs caused by these cells. As shown in Figure 6C, the addition of anti-IFN- mAb to coculture did not inhibit the activation of DCs by T cells. DCs matured in the presence of T cells from IFN--/- mice stimulated allogeneic T cells to the same degree as DCs from wt mice (Figure 6C). Similar results were observed with anti-TNF- mAb (data not shown). The percentage of DCs expressing CD40 was increased when cocultured with T cells. CD40L is a costimulatory molecule that is upregulated on CD4+ T cells after exposure to antigen and plays a critical role in DC activation.37-40 However, mAb blockade of CD40L did not prevent DC activation (Figure 6e). Furthermore, DCs from CD40-/- mice were activated to the same degree as DCs from wt mice, confirming that CD40-CD40L interactions are not required for T-cell-mediated DC activation (Figure 6D).
We next determined whether cell-to-cell contact was necessary for the T-cell activation of DCs. DCs were cultured for 36 hours either in direct contact with T cells or separated by a 0.2 μ transwell membrane and were then used as stimulators of allogeneic T cells. Physical separation of DCs and T cells in the transwells completely abrogated DC activation (Figure 6e), demonstrating the requirement for cell-to-cell contact.
Discussion
The localization of T cells within GVHD target tissues and their ability to modulate immune responses suggested that these cells might play a role in the pathogenesis of GVHD. In this study we demonstrate that the absence of T cells in allogeneic BM transplant recipients caused significantly less severe acute GVHD as determined by all measurable parameters in several different donor/recipient strain combinations. This finding was also confirmed by depleting T cells in wt recipients by in vivo administration of mAb against TCRs. Reconstitution of host T cells to -/- recipients increased lethal GVHD. Coculture of T cells with DCs enhanced their allostimulatory capacity and increased the proliferation of allogeneic responder T cells in vitro and in vivo. This enhancement was independent of IFN-, TNF-, and CD40L, but depended on cell-to-cell contact. These data demonstrate that host T cells play a critical role in the pathogenesis of GVHD by amplifying the stimulatory function of host DCs.
Donor T cells' interactions with host DCs are essential for the induction of GVHD.1-3 DCs represent a pool of professional APCs whose maturation/activation state is critical to their function. The chemo-radiotherapy used to condition BM transplant recipients induces the secretion of inflammatory cytokine such as IL-1, TNF-, and IL-6 from damaged host tissues,41 and allows LPS to enter into the systemic circulation.42 Such "danger signals" are critical for the activation of host DCs,43 but Shlomchik44 has reported that recipients deficient in TNFR1/R2, IL-1R, CD40, and Toll-like receptor 4 (essential for LPS-induced activation of DCs) developed GVHD comparable to that seen in wild-type controls. These experiments exclude a role for each of these individual pathways in the activation of host DCs and suggest that either together or through other pathways and/or cell interactions might contribute to the process. Several cell types such as macrophages,5 NK cells,45 CD4+ T,37-40 and CD8+ T46 have been shown to provide maturation and activation signals for DCs.
Our data demonstrate that murine T cells promote DC activation in a cell-to-cell contact-dependent manner. Leslie et al32 have also demonstrated that human T cells promote DC maturation in a cell-to-cell contact-dependent manner that is restricted by CD1c. Mice do not express CD1c but do express only CD1d, which presents glycolipid antigens to NKT cells.47 Recently, Hashimoto et al48 have demonstrated that absence of CD1d in BM transplant recipients results in a loss of protection from GVHD that is mediated by NKT cells. Thus, while our data demonstrated that T cells exacerbate GVHD, NKT cells can reduce GVHD, suggesting that in mice CD1 probably regulates GVHD through its interaction with NKT cells.
Other studies have focused on the role of T cells in the pathogenesis of GVHD.15-21 One study reported that a clonal population of T cells can induce lethal GVHD,17 whereas Drobyski and Majewski20 and Drobyski et al21 demonstrated that donor T cells alone do not cause acute GVHD. Coadministration of T cells and T cells exacerbated GVHD when compared with T cells alone, suggesting that donor T cells could contribute to GVHD but required the presence of T cells.18 However, the role played by host T cells in the pathogenesis of allogeneic GVHD has not been defined. Sakai et al27 reported that allogeneic BMT recipients depleted of T cells by in vivo administration of mAb against TCRs showed less intestinal damage, but the mechanisms for this phenomenon were not explored. We demonstrate that host T cells amplify systemic GVHD in irradiated GVHD models. Our data suggest that they might function as the "danger signals" and activate the DCs, thus enhancing their allostimulatory capacity.
Although T cells are a minor subpopulation of lymphocytes within lymph nodes and the spleen, they predominate in the epithelium of several principal target organs of GVHD, such as the skin, intestinal tract, and lung.49 T cells respond to infected or transformed epithelial cells and may therefore function as sensors of danger signals that can activate an immune response.43 Recent evidence suggests that IeLs express several chemokines such as macrophage inflammatory protein 1 (MIP1), MIP1, Regulated or Activation, Normal-T cell expressed and Secreted (RANTeS), and lymphotactin, and therefore play an important role in the rapid recruitment of CCR5+ cells into these epithelial tissues, helping to explain the target organ distribution of GVHD.50,51
T cells may also affect events in the gastrointestinal (GI) tract through their production of keratinocyte growth factor (KGF).52 KGF administration can reduce the severity of GVHD, particularly in the GI tract, and GVHD mortality.53,54 An absence of T cells would be expected to result in reduced endogenous KGF production and enhanced GI tract damage. To examine this possibility we compared the degree of tissue damage on day 7 after lethal irradiation and found no significant difference between wild-type and -/- mice (data not shown). Mice devoid of T cells showed decreased histopathologic damage from GVHD after allogeneic BMT (Figure 3), suggesting that whatever potential benefit T cells might have in GVHD by the secretion of KGF and subsequent promotion of tissue repair, it is overshadowed by the activation of host APCs and a greater donor response of allogeneic T cells.
In summary, we demonstrate a novel role for host T cells in the amplification of GVHD through the activation of host APCs, suggesting that elimination of host T cells might be a novel therapeutic strategy to reduce GVHD.
Footnotes
Prepublished online as Blood First edition Paper, March 29, 2005; DOI 10.1182/blood-2004-10-4087.
Supported by National Institutes of Health grants K08 AI052 863-01 (P.R.), DK 02 958 (C.L.), and CA 49 542 (J.L.M.F.). J.L.M.F. is a Doris Duke Distinguished Clinical Scientist.
An Inside Blood analysis of this article appears in the front of this issue.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
References
Zhang Y, Louboutin JP, Zhu J, Rivera AJ, emerson SG. Preterminal host dendritic cells in irradiated mice prime CD8+ T cell-mediated acute graft-versus-host disease. J Clin Invest. 2002;109: 1335-1344.
Shlomchik WD, Couzens MS, Tang CB, et al. Prevention of graft versus host disease by inactivation of host antigen-presenting cells. Science. 1999;285: 412-415.
Duffner UA, Maeda Y, Cooke KR, et al. Host dendritic cells alone are sufficient to initiate acute graft-versus-host disease. J Immunol. 2004;172: 7393-7398.
Mowat AM, Viney JL. The anatomical basis of intestinal immunity. Immunol Rev. 1997;156: 145-166.
Nestel FP, Price KS, Seemayer TA, Lapp WS. Macrophage priming and lipopolysaccharide-triggered release of tumor necrosis factor alpha during graft-versus-host disease. J exp Med. 1992;175: 405-413.
Ruggeri L, Capanni M, Urbani e, et al. effectiveness of donor natural killer cell alloreactivity in mismatched hematopoietic transplants. Science. 2002;295: 2097-2100.
Bryson JS, Flanagan DL. Role of natural killer cells in the development of graft-versus-host disease. J Hematother Stem Cell Res. 2000;9: 307-316.
Zeng D, Lewis D, Dejbakhsh-Jones S, et al. Bone marrow NK1.1(-) and NK1.1(+) T cells reciprocally regulate acute graft versus host disease. J exp Med. 1999;189: 1073-1081.
Chien YH, Hampl J. Antigen-recognition properties of murine gamma delta T cells. Springer Semin Immunopathol. 2000;22: 239-250.
Ferrick DA, Schrenzel MD, Mulvania T, Hsieh B, Ferlin WG, Lepper H. Differential production of interferon-gamma and interleukin-4 in response to Th1- and Th2-stimulating pathogens by gamma delta T cells in vivo. Nature. 1995;373: 255-257.
Jones-Carson J, Vazquez-Torres A, van der Heyde HC, Warner T, Wagner RD, Balish e. Gamma delta T cell-induced nitric oxide production enhances resistance to mucosal candidiasis. Nat Med. 1995;1: 552-557.
Skeen MJ, Ziegler HK. Activation of gamma delta T cells for production of IFN-gamma is mediated by bacteria via macrophage-derived cytokines IL-1 and IL-12. J Immunol. 1995;154: 5832-5841.
Nishimura H, emoto M, Hiromatsu K, et al. The role of gamma delta T cells in priming macrophages to produce tumor necrosis factor-alpha. eur J Immunol. 1995;25: 1465-1468.
Ladel CH, Blum C, Kaufmann SH. Control of natural killer cell-mediated innate resistance against the intracellular pathogen Listeria monocytogenes by gamma/delta T lymphocytes. Infect Immun. 1996;64: 1744-1749.
ellison CA, MacDonald GC, Rector eS, Gartner JG. Gamma delta T cells in the pathobiology of murine acute graft-versus-host disease: evidence that gamma delta T cells mediate natural killer-like cytotoxicity in the host and that elimination of these cells from donors significantly reduces mortality. J Immunol. 1995;155: 4189-4198.
Huang Y, Cramer De, Ray MB, Chilton PM, Que X, Ildstad ST. The role of alphabeta- and gammadelta-T cells in allogenic donor marrow on engraftment, chimerism, and graft-versus-host disease. Transplantation. 2001;72: 1907-1914.
Blazar BR, Taylor PA, Panoskaltsis-Mortari A, Barrett TA, Bluestone JA, Vallera DA. Lethal murine graft-versus-host disease induced by donor gamma/delta expressing T cells with specificity for host nonclassical major histocompatibility complex class Ib antigens. Blood. 1996;87: 827-837.
Drobyski WR, Vodanovic-Jankovic S, Klein J. Adoptively transferred gamma delta T cells indirectly regulate murine graft-versus-host reactivity following donor leukocyte infusion therapy in mice. J Immunol. 2000;165: 1634-1640.
Tsuji S, Char D, Bucy RP, Simonsen M, Chen CH, Cooper MD. Gamma delta T cells are secondary participants in acute graft-versus-host reactions initiated by CD4+ alpha beta T cells. eur J Immunol. 1996;26: 420-427.
Drobyski WR, Majewski D. Donor gamma delta T lymphocytes promote allogeneic engraftment across the major histocompatibility barrier in mice. Blood. 1997;89: 1100-1109.
Drobyski WR, Majewski D, Hanson G. Graft-facilitating doses of ex vivo activated gammadelta T cells do not cause lethal murine graft-vs-host disease. Biol Blood Marrow Transplant. 1999;5: 222-230.
Mombaerts P, Clarke AR, Rudnicki MA, et al. Mutations in T-cell antigen receptor genes alpha and beta block thymocyte development at different stages. Nature. 1992;360: 225-231.
Mombaerts P, Arnoldi J, Russ F, Tonegawa S, Kaufmann SH. Different roles of alpha beta and gamma delta T cells in immunity against an intracellular bacterial pathogen. Nature. 1993;365: 53-56.
Cooke KR, Kobzik L, Martin TR, et al. An experimental model of idiopathic pneumonia syndrome after bone marrow transplantation, I: the roles of minor H antigens and endotoxin. Blood. 1996;88: 3230-3239.
Cooke KR, Hill GR, Crawford JM, et al. Tumor necrosis factor-alpha production to lipopolysaccharide stimulation by donor cells predicts the severity of experimental acute graft-versus-host disease. J Clin Invest. 1998;102: 1882-1891.
Teshima T, Mach N, Hill GR, et al. Tumor cell vaccine elicits potent antitumor immunity after allogeneic T-cell-depleted bone marrow transplantation. Cancer Res. 2001;61: 162-171.
Sakai T, Ohara-Inagaki K, Tsuzuki T, Yoshikai Y. Host intestinal intraepithelial gamma delta T lymphocytes present during acute graft-versus-host disease in mice may contribute to the development of enteropathy. eur J Immunol. 1995;25: 87-91.
Wang T, Scully e, Yin Z, et al. IFN-gamma-producing gamma delta T cells help control murine West Nile virus infection. J Immunol. 2003;171: 2524-2531.
Hill GR, Crawford JM, Cooke KR, Brinson YS, Pan L, Ferrara JL. Total body irradiation and acute graft-versus-host disease: the role of gastrointestinal damage and inflammatory cytokines. Blood. 1997;90: 3204-3213.
Lutz MB, Kukutsch N, Ogilvie AL, et al. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J Immunol Methods. 1999;223: 77-92.
Duffner U, Lu B, Hildebrandt GC, et al. Role of CXCR3-induced donor T-cell migration in acute GVHD. exp Hematol. 2003;31: 897-902.
Leslie DS, Vincent MS, Spada FM, et al. CD1-mediated gamma/delta T cell maturation of dendritic cells. J exp Med. 2002;196: 1575-1584.
Fields RC, Shimizu K, Mule JJ. Murine dendritic cells pulsed with whole tumor lysates mediate potent antitumor immune responses in vitro and in vivo. Proc Natl Acad Sci U S A. 1998;95: 9482-9487.
Hill GR, Ferrara JL. The primacy of the gastrointestinal tract as a target organ of acute graft-versus-host disease: rationale for the use of cytokine shields in allogeneic bone marrow transplantation. Blood. 2000;95: 2754-2759.
Moore TA, Moore BB, Newstead MW, Standiford TJ. Gamma delta-T cells are critical for survival and early proinflammatory cytokine gene expression during murine Klebsiella pneumonia. J Immunol. 2000;165: 2643-2650.
Gao Y, Yang W, Pan M, et al. Gamma delta T cells provide an early source of interferon gamma in tumor immunity. J exp Med. 2003;198: 433-442.
Ridge JP, Di Rosa F, Matzinger P. A conditioned dendritic cell can be a temporal bridge between a CD4+ T-helper and a T-killer cell. Nature. 1998;393: 474-478.
Bennett SR, Carbone FR, Karamalis F, Flavell RA, Miller JF, Heath WR. Help for cytotoxic-T-cell responses is mediated by CD40 signalling. Nature. 1998;393: 478-480.
Schoenberger SP, Toes Re, van der Voort eI, Offringa R, Melief CJ. T-cell help for cytotoxic T lymphocytes is mediated by CD40-CD40L interactions. Nature. 1998;393: 480-483.
Cella M, Scheidegger D, Palmer-Lehmann K, Lane P, Lanzavecchia A, Alber G. Ligation of CD40 on dendritic cells triggers production of high levels of interleukin-12 and enhances T cell stimulatory capacity: T-T help via APC activation. J exp Med. 1996;184: 747-752.
Xun CQ, Thompson JS, Jennings CD, Brown SA, Widmer MB. effect of total body irradiation, busul-fan-cyclophosphamide, or cyclophosphamide conditioning on inflammatory cytokine release and development of acute and chronic graft-versus-host disease in H-2-incompatible transplanted SCID mice. Blood. 1994;83: 2360-2367.
Ferrara JL, Cooke KR, Teshima T. The pathophysiology of acute graft-versus-host disease. Int J Hematol. 2003;78: 181-187.
Gallucci S, Matzinger P. Danger signals: SOS to the immune system. Curr Opin Immunol. 2001;13: 114-119.
Shlomchik WD. Antigen presentation in graft-vs-host disease. exp Hematol. 2003;31: 1187-1197.
Zitvogel L. Dendritic and natural killer cells cooperate in the control/switch of innate immunity. J exp Med. 2002;195: F9-14.
Ruedl C, Kopf M, Bachmann MF. CD8(+) T cells mediate CD40-independent maturation of dendritic cells in vivo. J exp Med. 1999;189: 1875-1884.
Gumperz Je, Brenner MB. CD1-specific T cells in microbial immunity. Curr Opin Immunol. 2001;13: 471-478.
Hashimoto D, Asakura S, Miyake S, et al. Host NKT cells promote Th2 polarization of donor T cells and regulate acute GVHD after experimental BMT via a STAT6-dependent mechanism. Blood. 2003;102: 191A-192A.
Augustin A, Kubo RT, Sim GK. Resident pulmonary lymphocytes expressing the gamma/delta T-cell receptor. Nature. 1989;340: 239-241.
Shires J, Theodoridis e, Hayday AC. Biological insights into TCRgammadelta+ and TCRalpha-beta+ intraepithelial lymphocytes provided by serial analysis of gene expression (SAGe). Immunity. 2001;15: 419-434.
Boismenu R, Feng L, Xia YY, Chang JC, Havran WL. Chemokine expression by intraepithelial gamma delta T cells: implications for the recruitment of inflammatory cells to damaged epithelia. J Immunol. 1996;157: 985-992.
Boismenu R, Havran WL. Modulation of epithelial cell growth by intraepithelial gamma delta T cells. Science. 1994;266: 1253-1255.
Panoskaltsis-Mortari A, Lacey DL, Vallera DA, Blazar BR. Keratinocyte growth factor administered before conditioning ameliorates graft-versus-host disease after allogeneic bone marrow transplantation in mice. Blood. 1998;92: 3960-3967.
Krijanovski OI, Hill GR, Cooke KR, et al. Keratinocyte growth factor separates graft-versus-leukemia effects from graft-versus-host disease. Blood. 1999;94: 825-831.(Yoshinobu Maeda, Pavan Re)