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Departments of
*
Medicine,
Pediatric Immunology, and
Pathology, University of Massachusetts Medical School, Worcester, MA 01605; and
§
The Jackson Laboratory, Bar Harbor, ME 04609
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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EBV-related LPD that develop in C.B-17-scid mice are similar to those that develop in immunocompromised humans (5, 7, 8, 9, 10, 11, 12, 13). The human immune response to EBV is characterized by the development of cytotoxic CD8+ T cells (14, 15, 16), but the causal role of these cytotoxic CD8+ T cells in protecting the host from EBV-related LPD is unknown. Studies of the human immune response to EBV-infected cells have been hindered by the lack of a suitable animal model that demonstrates immune-mediated regulation of EBV-related LPD in vivo.
We have shown that the NOD/LtSz-Prkdcscid/Prkdcscid (NOD-scid) mouse is an improved immunodeficient host for human hematolymphoid cell engraftment (17, 18, 19). This strain of mouse supports levels of human cell engraftment that are 5- to 10-fold greater than those obtained in C.B-17-scid mice (3, 17). We have also shown that NOD-scid ß2-microglobulin-null (NOD-scid-B2mnull) mice support even higher levels of engraftment, and that this engraftment is associated with increased levels of human CD4+ T cells (20). The increased level of engraftment in NOD-scid-B2mnull mice has been postulated to be the result of an almost complete absence of NK cell activity (20). The engrafted cells in both strains of NOD-scid mice, however, continue to express an activated phenotype based on expression of cell surface markers that is similar to that observed in Hu-PBL-C.B-17-scid mice (3). Surprisingly, despite the increased engraftment of human cells, the development of EBV-related LPD has not been reported in NOD-scid or NOD-scid-B2mnull mice (3).
In the present report we identify the basis for the increased engraftment of human cells in Hu-PBL-NOD-scid mice and document a low frequency of EBV-related LPD in these mice. We demonstrate that the model is suitable for the in vivo analysis of immunoregulatory interactions between human CD4+ and CD8+ cells, and that human CD8+ cells regulate the in vivo expression of EBV-related LPD.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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NOD/LtSz-Prkdcscid/Prkdcscid (NOD-scid), NOD/LtSz-Prkdcscid/Prkdcscid-B2mnull (NOD-scid-B2mnull), and C.B-17-Prkdcscid/Prkdcscid (C.B-17-scid) mice were obtained from colonies maintained by LDS at The Jackson Laboratory (Bar Harbor, ME). All animals were certified to be free of Sendai virus, pneumonia virus of mice, murine hepatitis virus, minute virus of mice, ectromelia, lactate dehydrogenase (LDH)-elevating virus, GD7 virus, Reo-3 virus, mouse adenovirus, lymphocytic choriomeningitis virus, polyoma, Mycoplasma pulmonis, and Encephalitozoon cuniculi. All animals were housed under specific pathogen-free conditions in microisolator cages and given ad libitum access to autoclaved food. All immunodeficient mice were maintained on an alternate schedule of autoclaved water and a solution of sulfamethoxazole-trimethoprim (Goldline Laboratories, Ft. Lauderdale, FL) (20). All animals were maintained in accordance with recommendations in the Guide for the Care and Use of Laboratory Animals (Institute of Laboratory Animal Resources, National Research Council, National Academy of Sciences, 1996) and the guidelines of the institutional animal care and use committee of the University of Massachusetts Medical School.
Antibodies
Anti-mouse CD45 and anti-human CD4, CD8, CD25, CD45, CD45R0,
and HLA-DR flurochrome-conjugated mAb for flow cytometry and purified
sodium azide-free anti-human CD3 and CD28 mAb for in vitro
proliferation assays were obtained from PharMingen (San Diego, CA).
mAbs specific for human CD3, CD20, and Ig
and Ig
chain purchased
from Becton Dickinson (Mountain View, CA) were used for
immunoperoxidase staining. Ascites was generated from hybridomas
TM-ß1 (anti-mouse CD122, IL-2R ß-chain, rat IgG2b
(21), gift from T. Tanaka, Osaka University Medical
Center), OKT4 (anti-human CD4, mouse IgG2b; American Type Culture
Collection, Manassas, VA), OKT8 (anti-human CD8, mouse IgG2a;
American Type Culture Collection), and B73.1 (anti-human CD16,
mouse IgG1, gift from Dr. Giorgio Trinchieri, The Wistar Institute,
Philadelphia, PA). The hybridoma Ab 24-31 (anti-human CD154, mouse
IgG1) was a gift from Dr. Randolph Noelle, Dartmouth University
(Lebanon, NH). The ascites for in vivo depletion studies was collected
and pooled, the fibrin clot was removed by filtration through nylon
wool, and aliquots were frozen until used. mAb concentrations were
quantified using anti-mouse IgG radial immunodiffusion plates (The
Binding Site, San Diego, CA). The mAbs were diluted to a concentration
of 1 mg/ml in PBS immediately before injection.
Cell preparations
Splenic single cell suspensions were prepared from Hu-PBL-scid mice by teasing spleens between frosted microscope slides. RBC were lysed in hypotonic ammonium chloride lysing buffer and rinsed twice in PBS supplemented with 5% FBS (HyClone, Logan UT), and the number of viable cells was determined by exclusion of trypan blue using a hemocytometer. Peritoneal cavity mononuclear cells were recovered from Hu-PBL-scid mice at necropsy by lavaging the peritoneal cavity with 5 ml of RPMI 1640 warmed to 37°C.
Purified populations of human CD4+ and CD8+ peripheral blood T cells were prepared by positive selection using immunomagnetic beads (Dynabeads M-450, Dynal, Oslo, Norway). Cells were detached from the beads using the Detach-a-Bead reagent (Dynal) according to the manufacturer’s instructions.
B lymphoblastoid cell lines (B-LCL) were initiated from donor peripheral blood mononuclear cells by induction with supernatant from the EBV-producing cell line B95.8 as previously described (22). B-LCL lines were maintained in culture for use as targets or stimulators for EBV-specific T cell analyses.
To generate EBV-specific or allo-specific CTLs, standard protocols were used. Briefly, NK-depleted PBMC were cultured in complete medium with mitomycin C-treated autologous or allogeneic B-LCLs, respectively. Cultures were restimulated weekly with mitomycin C-treated B-LCL and fed twice weekly with fresh complete medium containing 20 U/ml recombinant human IL-2. Cell lines were tested for phenotype and cytolytic activity after 3–4 wk in culture.
Relative cytotoxic activity of EBV-specific and allo-specific CTLs was determined by calculating lytic units based on cytolytic activity in a 51Cr release assay (23). The allo-specific CTLs were cytotoxic to allogeneic B-LCLs, but displayed no cytotoxic activity against autologous B-LCLs (data not shown).
Experimental treatments
Establishment of Hu-PBL-scid mice.
Human PBMC were prepared from normal platelet-pheresis donors as
previously described (18). Unless otherwise noted, mice
were injected i.p. with 20 x 106 human PBMC
in RPMI within 3 h of recovery following purification of human
mononuclear cells by Ficoll-Hypaque density gradient centrifugation. In
certain studies of EBV-related LPD development, 50–60 x
106 human PBMC were injected i.p. Analysis of
engrafted mice was performed 4 wk after injection of PBMC unless
otherwise noted. For ease of presentation, we have provided results
from a single donor in Table I
. With the
single exception of the histology photomicrograph representative of
multiple animals from multiple donors, we show pooled results from
multiple donors in all other tables and figures.
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Histology and immunohistochemistry
Tissues were recovered from mice at necropsy, fixed in 10%
buffered formalin, and embedded in paraffin. Sections 4 µm thick were
cut. For routine histology, sections were stained with hematoxylin and
eosin. The sections were interpreted by B.A.W. without knowledge of the
experimental design. For immunohistochemistry, sections were heated at
60°C for 30 min, then deparaffinized and hydrated through a series of
xylene and alcohol baths before staining. The slides were microwaved in
a proprietary citrate-buffered Ag retrieval solution (BioTek Solutions,
Santa Barbara, CA) for 5 min in an 800-W microwave oven. After
replenishment of this solution the slides were microwaved for another 5
min, then allowed to cool for 20 min. Immunohistochemical staining was
performed with mAb specific for human CD3, CD20, Ig, and Ig
chains using a standard avidin/biotin complex method as implemented on
a Techmate 1000 (BioTek) automated immunostainer. The sections were
counterstained with hematoxylin. To detect EBV early RNA (EBER) by in
situ hybridization, sections were deparaffinized and washed in 2x
sodium chloride/sodium citrate buffer. The sections were hybridized
with a biotinylated EBER oligonucleotide probe at 37°C for 18 h.
After washing in sodium chloride/sodium citrate buffer, the bound probe
was detected using the avidin/biotin complex method.
Quantification of human Ig levels
Blood was collected from the retro-orbital plexus of individual engrafted mice, and levels of human Ig in sera were determined by nephlometry using a Beckman ARRAY 360 CD Serology System (Beckman Coulter, Fullerton, CA).
Mixed lymphocyte cultures
Murine stimulator cells were prepared by teasing spleens recovered from NOD-scid or NOD-scid-B2mnull mice between frosted microscope slides to obtain single-cell suspensions. RBC were lysed with hypotonic ammonium chloride lysing solution, and the nucleated cells were washed in RPMI supplemented with 10% FBS, 100 U/ml penicillin G, 100 µg/ml streptomycin, and 0.29 mg/ml L-glutamine (complete medium); counted; and suspended in complete medium at 8 x 106 cells/ml. The stimulator cells were then exposed to 2000 rad 137Cs radiation using a Gammacell 40 (Atomic Energy of Canada, Ottawa, Canada). Murine stimulator cells (8 x 105 in 100 µl) in complete medium were added to each well of 96-well flat-bottom tissue culture plates (Falcon, Becton Dickinson Labware, Lincoln Park, NJ). Responder human PBMC (2.5 x 105 in 100-µl aliquots) were added to triplicate wells, and the mixture was cultured for 6 days at 37°C in a humidified atmosphere of 95% air in 5% CO2. For the last 16 h of culture, the cells in some plates were incubated with 1 µCi of [3H]thymidine (New England Nuclear, Boston, MA). At the end of culture, the pulsed cells were harvested onto filter mats using an Quadra96 harvester (Tomtec, Hamden, CT), and total incorporated radioactivity was determined using a Wallac 1450 Microbeta scintillation counter (EG&G Wallac, Turku, Finland). Cells from separate wells that were treated identically but not incubated in the presence of [3H]thymidine were analyzed for activation marker expression by flow cytometry.
Ex vivo proliferation cultures
Pools of Hu-PBL-NOD-scid and Hu-PBL-NOD-scid-B2mnull splenocytes were washed, counted, and suspended in complete medium at a concentration of 2.5 x 105 cells in 200 µl. Triplicate cultures were stimulated with 1 µg of anti-human CD3 mAb and 0.5 µg of anti-human CD28 mAb (PharMingen) that was immobilized on the culture plate. In some wells, 100 U/ml recombinant human IL-2 (Endogen, Cambridge, MA) was added. Cells were cultured for 6 days at 37°C in a humidified atmosphere of 95% air and 5% CO2 and incubated for the last 16 h of culture in the presence of 1 µCi of [3H]thymidine. The cultures were harvested, and incorporated radioactivity was quantified as described above.
Flow cytometry
Spleen or peritoneal cavity cells recovered from Hu-PBL-scid mice were concentrated by centrifugation and washed in PBS plus 5% FBS, aliquoted into microtiter plates, and prepared for three-color flow cytometric analysis by incubation with PE-anti-murine-CD45, anti-human CD4 or anti-human CD8 conjugated to PE-Cy5, and FITC-conjugated anti-human CD25, anti-human CD45R0, or anti-human HLA-DR (17, 18, 19). Cells were incubated in the presence of 1 µg of Ab/106 cells for 20 min at 4°C, washed three times in PBS plus 5% FBS, and fixed in 1% paraformaldehyde in PBS plus 0.02% NaN3 for analysis.
At least 15,000 events were acquired on Becton Dickinson FACS equipment (Becton Dickinson, San Jose, CA), and the data were analyzed with WinMDI software (J. Trotter, The Scripps Research Institute, La Jolla, CA). In all analyses, murine CD45+ cells labeled with anti-Ly5 mAb were excluded from further analysis. On the remaining cells, a two-color histogram of CD4 or CD8 vs an activation or proliferation marker (forward scatter, CD25, CD45R0, or HLA-DR) was used to determine the percentage of double-positive cells. Percentages are the mean ± SD of CD4+ or CD8+ cells that were also positive for the expression of an activation marker (CD25, CD45R0, or HLA-DR). The percentage of large cells was determined by increases in the forward light scattering properties compared with freshly isolated, nonstimulated CD4+ or CD8+ cells. Isotype Ab controls were used to assist in the determination of negative and positive populations.
Analysis of cell proliferation using 5-(and-6)-carboxyfluorescein succinimidyl esters (CFSE)-labeled cells
Human PBMC were labeled with CFSE (Molecular Probes, Eugene OR) as described previously (24). Labeled cells (20 x 106) were injected into the peritoneal cavity of NOD-scid mice, NOD-scid-B2mnull mice, and NOD-scid mice treated previously with anti-mouse CD122 mAb. The cells were recovered 6 days later by lavaging the peritoneal cavity with 5 ml of RPMI 1640 warmed to 37°C. Recovered cells were suspended in PBS plus 5% FBS, aliquoted into 96-well microtiter plates and labeled with anti-murine CD45-PE and either anti-human-CD4 or anti-human-CD8 mAb conjugated to PE-Cy5. Labeled cells were washed twice in PBS plus 5% FBS, fixed in 1% paraformaldehyde, and analyzed by flow cytometry.
Statistics
In experiments on development of EBV-related LPD, p
values were calculated by 
2 analysis using
StatView software (Abacus Concepts, Berkley, CA). Median survival times
(MST) of Hu-PBL-scid mice were calculated using the method
of Kaplan and Meier (25). The equality of survival time
distributions for animals in different treatment groups was tested
using the log rank statistic (25). For analysis of
proliferation using CFSE dye labeling (Table II) and for analysis of human cells in
the spleen of scid mice in Table IV, comparisons of means
used one-way ANOVA (26) and the Bonferroni test for a
posteriori contrasts (27). For comparison between multiple
groups in the remaining experiments, we used the Bonferroni adjusted
unpaired t test (27).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Human PBLs in C.B-17-scid mice have been reported to
have an activated phenotype, but are functionally anergic
(28, 29, 30). To determine the kinetics of progression of
engrafted human cells from that of a naive to an activated to an
anergic phenotype, peritoneal cavity and spleen cells from separate
cohorts of Hu-PBL-NOD-scid and
Hu-PBL-NOD-scid-B2mnull mice
established from a single PBMC donor were analyzed at 1, 2, and 3 wk
after engraftment. The phenotype of peritoneal cavity cells 1 wk after
engraftment resembled that of recently activated cells (Table I
). By 2
and 3 wk after engraftment, however, the phenotypic characteristics of
human cells in the peritoneal cavity resembled those observed in the
spleen at 2 and 3 wk (Table I). The percentage of
CD25+ T cells in both tissues progressively
declined, and the proportion of cells exhibiting characteristics of
activated, CD25- T cells increased with similar
kinetics in both the peritoneal cavity and spleen. There was no
evidence of tissue compartmentalization of naive or activated human
cells between the peritoneal cavity and spleen at 2 and 3 wk after
engraftment, time points when direct comparisons were possible. The
phenotype of human cells in the spleen at 4 wk was similar to that
observed at 3 wk (data not shown).
To confirm that the activated CD25- human T
cells at 4 wk after engraftment exhibited functional characteristics
consistent with an anergic phenotype, we performed in vitro analyses.
Human cells recovered from the spleens of Hu-PBL-NOD-scid
and Hu-PBL-NOD-scid-B2mnull
mice that were incubated in the presence of plate-bound anti-human
CD3 and anti-human CD28 mAb did not proliferate (Fig. 1). Addition of recombinant human IL-2,
however, supported high levels of proliferation (Fig. 1). This
functional characteristic is consistent with that of an anergic
population described previously for human cells recovered from
PBMC-engrafted C.B-17-scid mice (2).
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NOD-scid-B2mnull mice engraft higher levels of human cells than do NOD-scid mice (20). This may result from differential expression of MHC class I on the murine stimulator cells, because the B2mnull allele prevents MHC class I expression. To quantify the in vivo proliferative activity of human CD4+ and CD8+ cells, we employed the intravital dye CFSE. This dye labels lymphoid cells (24), and the uniform dilution of dye in cells as they divide can be used to determine whether labeled cells have undergone cell division (31).
Human PBMC labeled with CFSE were injected into the peritoneal cavity
of NOD-scid and
NOD-scid-B2mnull mice and
recovered 1 wk after engraftment for flow cytometric analysis. The
percentage of CD8+ CFSE-labeled cells that had
undergone division in
NOD-scid-B2mnull mice was
significantly greater than that observed in NOD-scid mice
(Table II).
Activation of human T cells by NOD-scid and NOD-scid-B2mnull APC
We next analyzed the activation of human cells by
NOD-scid and
NOD-scid-B2mnull mice using in
vitro xenogeneic mixed lymphocyte cultures. There were no detectable
significant differences between the activation phenotype of human
CD4+ or CD8+ T cells
incubated with NOD-scid irradiated splenocytes compared with
that observed following stimulation with
NOD-scid-B2mnull irradiated
splenocytes (Table III). The
CD4+ and CD8+ T cell
subsets in both cultures exhibited a phenotypic pattern consistent with
activation, including comparable increases in forward light-scattering
properties and cell surface expression of CD25, CD45R0, and HLA-DR
(Table III).
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Human cell engraftment in NOD-scid mice is dependent on host NK cell activity
We next investigated whether the increased engraftment of human
cells in NOD-scid-B2mnull mice
was due to their severe deficiency in NK cell activity compared with
the relatively less severe deficiency of NK cells in
NOD-scid mice (20). To test this, murine NK
cells were depleted in vivo in a cohort of NOD-scid mice
using the anti-mouse CD122 mAb TM-ß1 that recognizes the mouse
IL-2R ß-chain expressed on NK cells (32). At 4 wk after
engraftment Hu-PBL-NOD-scid mice treated with anti-mouse
CD122 mAb had significantly higher percentages of human cells in their
spleens than did Hu-PBL-NOD-scid mice that did not receive
TM-ß1 mAb (Fig. 2). The absolute number
of human cells in the spleen was also elevated, with an average of
18 x 106 ± 4.5 x
106 cells in anti-mouse CD122 mAb-treated
hosts (mean ± SEM; n = 15) compared with 1.4
x 106 ± 0.8 x 106
cells in untreated hosts (n = 19; p <
0.003).
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Treatment with TM-ß1 anti-mouse CD122 mAb also enhanced the
proliferation of engrafted cells in NOD-scid mice by day 6
as measured by CFSE staining. The percentages of
CD4+ cells (79.5 ± 10.4%) and
CD8+ cells (77.4 ± 9.5%) that had
undergone cell division in mice treated with TM-ß1 mAb was
significantly higher than that observed in either NOD-scid
or NOD-scid-B2mnull mice
injected with PBMC from the same donor (Table II).
CD4+ cells and CD40-CD154 interactions are required for engraftment of human cells in scid mice
The host innate immune system clearly has a critical role in the
cell subset and number of human cells that engraft in
Hu-PBL-scid mice. We next studied the roles of the human
CD4+ and CD8+ T cell
subsets in regulating human cell engraftment in NOD-scid and
NOD-scid-B2mnull mice. To
prevent human CD4+ cell engraftment, we treated
Hu-PBL-NOD-scid and
Hu-PBL-NOD-scid-B2mnull mice
with a depleting anti-human CD4 mAb. Although both
CD4+ and CD8+ human cells
readily engrafted in untreated recipients, specific depletion of only
CD4+ cells led to the ablation of both human
CD4+ and CD8+
cell engraftment (Table IV). As shown
previously (33), human CD4+ and
CD8+ cell engraftment was also inhibited by
treatment with anti-human CD154 mAb, a costimulation molecule
expressed predominately on activated CD4+, but
not CD8+, T cells (Table IV).
CD8 cells regulate CD4 cell expansion in Hu-PBL-scid mice
To determine whether human CD8+ cells are
required for CD4+ cell engraftment, we treated a
cohort of Hu-PBL-NOD-scid and
Hu-PBL-NOD-scid-B2mnull mice
with a depleting anti-human CD8 mAb. Engraftment of total human
cells in the spleen at 4 wk, as determined by expression of human CD45,
was similar to that obtained in otherwise untreated
Hu-PBL-NOD-scid and
Hu-PBL-NOD-scid-B2mnull mice
(Table IV). However, the majority of the engrafted T cells were
CD4+; few to no human CD8+
cells were detected in the blood or spleen (Table IV).
Corresponding with increased engraftment of human CD4+ cells in the spleen, the levels of human Ig in Hu-PBL-NOD-scid mice following treatment with anti-human CD8 mAb were significantly increased (497.5 ± 280.6 µg/ml IgG and 151.4 ± 81.6 µg/ml IgM; n = 7) compared with those observed in control Hu-PBL-NOD-scid mice (80.5 ± 33.7 µg/ml IgG (p < 0.02); 21.0 ± 9.0 µg/ml IgM (p < 0.02); n = 13).
Depletion of CD8+ cells increases the development of EBV-related lymphoproliferative disorders in Hu-PBL-NOD-scid mice
Injection of high numbers of PBMC (
50 x
106) obtained from EBV-seropositive donors
frequently leads to the outgrowth of EBV-related LPD in
Hu-PBL-C.B-17-scid mice (6, 11). Despite
increased levels in the engraftment of human cells in
Hu-PBL-NOD-scid mice following the injection of 20 x
106 human PBMC, we observed that few mice
exhibited EBV-related LPD that were detectable by necropsy at 4–6 wk.
To quantify this, we injected separate cohorts of NOD-scid
and C.B-17-scid mice with50–60 x
106 PBMC obtained from two different
EBV-seropositive donors. In Hu-PBL-NOD-scid mice,
EBV-related LPD were observed at the 16 wk necropsy in 4 of 12 mice
(33%) injected with PBMC from donor 1 and in 25 of 46 (54%) of mice
injected with PBMC from donor 2. In contrast, injection of PBMC into
C.B-17-scid mice led to the development of EBV-related LPD
in 35 of 38 (92%; p < 0.002) and 27 of 32 (84%;
p < 0.004), respectively, of the recipients compared
with PBMC from the same donors in NOD-scid mice.
Hu-PBL-NOD-scid mice treated with anti-human CD8 mAb had increased levels of human CD4+ cells and increased levels of circulating human Ig. To investigate the basis for these observations, we performed a morphologic analysis of spleen, lymph node, liver, and kidney obtained from cohorts of untreated and CD8-depleted Hu-PBL-NOD-scid mice 4 wk after engraftment.
No detectable evidence of EBV-related LPD at necropsy 4 wk after
engraftment was observed upon analysis of tissues removed from
Hu-PBL-NOD-scid mice. Some animals did show moderate
expansion of the white pulp and lymph node paracortex. Such animals
usually did not show any appreciable interstitial infiltrate of the
kidney and showed only moderate periportal hepatic infiltrates. Other
animals showed massive lymphoid expansion in spleen, lymph node, liver,
and kidney. The lymph nodes or spleens never evidenced development of
germinal centers. The lymphoid infiltrates were diffuse and composed
predominantly of large activated lymphoid cells. Scattered plasma cells
were also noted. In situ hybridization studies were performed to detect
EBER, indicative of latent EBV infection. Four of the 10 otherwise
untreated Hu-PBL-NOD-scid mice had detectable EBER. We also
localized human CD3+,
CD20+, Ig+, and
Ig+ cells by immunohistochemistry. These
studies showed that in the EBER-negative animals the lymphoid cells
were composed predominantly of CD3+ cells.
CD20+ cells and plasma cells were few. When EBER
was detectable, CD20+ and plasma cells were
increased. In some of these animals, numerous
CD20+ cells and plasma cells that expressed Ig
and Ig chains were detected.
The morphologic features of the tissues examined from the CD8-depleted
animals were uniform. These animals (six of six; by
2 test, p < 0.01 vs
non-CD8-depleted animals) all shared detectable EBER-positive cells. In
some animals only a few EBER positive cells were detectable, while in
others many positive cells were noted (Fig. 3). Histologic examination showed a
marked proliferation of large transformed lymphoid cells (Fig. 3). The
infiltrates were quite polymorphous and were composed of a mixture of
lymphoblasts, immunoblasts, and plasma cells. Germinal centers were
never identified. Immunoperoxidase studies (Fig. 3, C and
D) generally showed both CD3+ T cells
and CD20+ B cells. In some tissues
CD20+ B cells and Ig+
and Ig+ plasma cells were numerous.
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We next confirmed that human CD8+ T cells
were the primary mediators of resistance in vivo to the development of
EBV-related LPD in Hu-PBL-NOD-scid mice. Purified
populations of CD4+ or CD8+
human T cells were adoptively transferred into NOD-scid mice
that had received 1) 50–60 x 106 PBMC from
the same original donor and 2) a single injection of anti-CD8 mAb
at 1–3 days after cell engraftment. Evidence for the development of
EBV-related LPD was apparent at necropsy 16 wk later in 29 of 58 (50%)
of the Hu-PBL-NOD-scid mice that received cells without
coinjection of anti-CD8 Ab (Fig. 4).
This rate was comparable to that observed in our initial study above.
CD8-depleted Hu-PBL-NOD-scid mice uniformly (20 of 20,
100%) developed EBV-related LPD by 16 wk after cell engraftment (Fig. 4). In contrast, i.p. injection of 5–10 x
106 CD8+ T cells from the
original PBMC donor on days 16 and 34 after initial PBMC injection
restored CD8+ T cell engraftment and reduced the
incidence of EBV-related LPD in CD8-depleted Hu-PBL-NOD-scid
mice from 100% to only 1 of 10 (10%). The transfer of 5–10 x
106 CD4+ T cells into
CD8-depleted Hu-PBL-NOD-scid mice on the same schedule was
unable to prevent the development of EBV-related LPD (7 of 9, 78%;
Fig. 4). The depletion of donor NK cells by injection of anti-human
CD16 mAb 1–3 days after PBMC engraftment had no effect on the
proportion of mice that developed EBV-related LPD by 16 wk (6 of 17,
35%; Fig. 4).
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We next determined whether EBV-specific CD8+ T cells could delay or prevent EBV-related LPD in Hu-PBL-C.B-17-scid mice that develop this disorder at high frequency. To address this, we engrafted C.B-17-scid mice with 50–60 x 106 human PBMC from two different EBV-seropositive donors. Engraftment of these cell numbers is known to lead to the development of EBV-related LPD in the majority of scid mice (4, 34). One to 3 days after PBMC injection and twice weekly thereafter, 100–200 lytic units of CTL specific for autologous EBV+ B-LCLs or allogeneic B-LCLs were injected i.p.
The cumulative survival of Hu-PBL-C.B-17-scid mice
receiving either EBV-specific CTL (MST, 81 days; range, 49–95 days;
n = 13) or allo-specific CTL (MST, 83 days; range,
62–89 days; n = 9) was significantly increased
compared with that of untreated mice (MST, 62 days; range, 45–89 days;
n = 23; p = 0.006 and p
= 0.0008, respectively). Both types of CTL immunotherapy led to a delay
in tumor outgrowth, but did not prevent tumors from developing. There
was no statistical difference in the survival of recipients of
EBV-specific and allo-specific CTLs (p = 0.26).
Surviving mice were sacrificed on day 89 (untreated and allo-specific
CTL-treated) or on day 95 (EBV-specific CTL-treated) and examined
visually for evidence of EBV-related LPD. One untreated mouse (4.3%),
three mice treated with EBV-specific CTL (23.1%), and two mice treated
with alloreactive CTL (22.2%) were free of tumors at the end of the
experiment, but these values were not statistically significant
(p = 0.19, by 2
analysis).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our previous studies have demonstrated that host strain characteristics influence the engraftment of human PBMC (17, 20). NOD-scid mice lack hemolytic complement, have defects in myeloid cells, and are relatively deficient in NK cell activity (19). NOD-scid-B2mnull mice retain all these innate immune deficiencies, but in addition are severely deficient in NK cell activity and lack murine MHC class I expression. NOD-scid-B2mnull mice support heightened levels of human T cell engraftment, and in contrast to that observed in Hu-PBL-NOD-scid mice, there was a normal ratio of CD4:CD8 human cells in Hu-PBL-NOD-scid-B2mnull mice (20). The basis of these differences in human cell engraftment could result from differences in host NK cell activity or from the absence of host MHC class I (20, 35, 36).
To dissect the mechanisms that drive human T cell engraftment in NOD-scid and NOD-scid-B2mnull mouse recipients, we evaluated the proliferation of human cells in the first week of engraftment in NOD-scid-B2mnull mice compared with that in NOD-scid mice. No differences in proliferation were observed in vitro in primary cultures following stimulation of fresh human PBMC with irradiated splenocytes from each strain. These results suggest that a deficiency in expression of MHC class I does not alter the direct proliferative stimulus of mouse splenocytes to human lymphocytes. Moreover, additional studies demonstrated that increased human T cell proliferation and a normal CD4:CD8 ratio could be obtained in NOD-scid mice following injection of TM-ß1 anti-mouse CD122 mAb. IL-2R ß-chain (CD122) is expressed on mouse NK cells, granulocytes, and, to a lesser extent, many other cell types (37, 38). Administration of TM-ß1 mAb to Hu-PBL-C.B-17-scid mice has been shown to increase human cell engraftment (36). Our present data show that human cell proliferation is increased in NOD-scid mice treated with anti-mouse CD122 mAb, and that this treatment leads to a CD4:CD8 ratio similar to that observed in NOD-scid-B2mnull mice. Our interpretation of these results is that the high engraftment of CD4+ cells and the normal CD4:CD8 ratio observed in Hu-PBL-NOD-scid-B2mnull mice are the result of their almost total absence of NK activity.
In Hu-PBL-NOD-scid mice, the progression of human cells from naive to activated to an anergic phenotype proceeds concurrently in the peritoneal cavity and the spleen. These data are consistent with previous reports of low levels of activation 1 wk after engraftment (39) and expression of an anergic phenotype of human cells in the spleen of mice by 4–6 wk after engraftment (2). These observations suggest that the activation status of human cells does not determine whether they will localize in the spleen or peritoneal cavity in the scid host. By 4 wk, the proliferation of human cells in the spleen of NOD-scid mice is low and correlates kinetically with their loss of CD25 expression and acquisition of an anergic phenotype. Although overall levels of human cell engraftment and human CD4+ T cell engraftment were increased in NOD-scid mice, the human cells still displayed functional and phenotypic defects in activation. These phenotypic characteristics are reported to be associated with an anergic phenotype (2).
A new observation reported here is the regulation of human cell engraftment by human CD4+ and CD8+ cells. No detectable engraftment is observed in the absence of human CD4+ cells. These results demonstrate that there is a requirement for CD4-derived helper factors, perhaps IL-2, for the expansion of CD8+ cells in vivo. This help is probably induced through expression of CD154 and its interaction with CD40 on APC. In other studies we have observed that expansion of TCR transgenic alloreactive CD8+ cells in vivo is absolutely dependent on CD4 helper factors that are independent of CD40-CD154 interaction (40). These earlier results are similar to those obtained in the present experiments. The expression of CD25 on activated human cells at 1 wk, but not at 4 wk, after engraftment also supports this possibility, as does restoration of proliferation in vitro following addition of recombinant human IL-2.
We also document that the regulation of CD4+ cell expansion by CD8+ cells is a potentially novel model system with which to study CD8 regulatory activity in vivo. The mechanism by which this regulation occurs is unknown, but in other regulatory systems CD95- or TNF-mediated apoptosis, TGF-ß, and T cell-NK cell interactions have all been implicated as mediators of suppression (41). In preliminary experiments we have obtained evidence that suppression of CD4+ cell engraftment may be mediated in part by elimination of activated CD4+ cells via CD95-CD95L (Fas-Fas ligand) cytotoxicity. Both CD4+ and CD8+ cells at 1 wk of engraftment express high levels of Fas, and preliminary in vitro experiments suggest that activated human CD4+ cells are more susceptible than activated human CD8+ cells to CD95L-mediated apoptosis (E. J. Wagar, unpublished observations).
The possible role of CD8+ cells as suppressor cells is consistent with several previous reports that have provided data on the in vivo function of CD8+ cells. These earlier studies have generated suggestive data that CD8+ cells (42, 43, 44) or anergic T cells can act as suppressor T cells (41, 45, 46). The mechanism of suppression has not been identified in these published studies.
Of additional interest is our documentation that Hu-PBL-NOD-scid mice are relatively resistant to the development of EBV-related LPD. Inhibition of EBV-related LPD in Hu-PBL-NOD-scid mice may be mediated by the increased levels of engraftment of human CD8+ cells (47, 48, 49). This interpretation is based on previous data obtained in humans (50, 51, 52, 53) and in scid mice bearing EBV-related LPDs (54). In humans, transfer of autologous EBV-specific CTL prevents or delays the development of EBV-related LPD following bone marrow (50, 51, 52) or solid organ transplantation (53). In mice bearing B-LCL tumors, autologous EBV-specific CTLs can induce the regression of coinjected or established B-LCL tumors in the host (54). Our data suggest that human CD8+ cells in NOD-scid mice remain functional and mediate the suppression of EBV-related LPD outgrowth. This inference was confirmed by data obtained in CD8-depleted Hu-PBL-NOD-scid mice and in CD8-depleted mice that were reconstituted with human CD8+ cells. Published data suggest that human EBV-specific CTLs can induce regression against established EBV-related LPD in scid mice (54). Our current data suggest further that human CD8+ cells also provide surveillance activity to prevent the development of EBV-related LPD in Hu-PBL-C.B-17-scid mice. This observation in consistent with the data in humans demonstrating that IL-2-activated CD8+ T cell immunotherapy can prevent post-transplant LPD (55). The requirement for EBV-specific CTL vs nonspecifically activated CD8+ T cells in surveillance is under investigation. Our studies also ruled out a major role for human NK cell control of EBV-related LPD.
We also observed that CD8+ cells regulated human CD4+ cell outgrowth. This may be due to a direct suppressive effect, as described above, or indirectly by prevention of EBV-related LPD outgrowth. The expanding EBV+ cells would provide high levels of human MHC class II and multiple costimulatory factors that are normally absent in Hu-PBL-NOD-scid mice. This, in turn, could stimulate human CD4+ cell proliferation. Finally, human CD8+ cells appeared to modulate the levels of circulating human Ig. This effect could be directly on Ig-producing B lymphocytes or a secondary effect due to regulating the expansion of EBV-infected cells. These data document the important role of CD8-mediated inhibition of B cell activation, and the complex balance between CD4 help and CD8 suppression on B cell function and lymphoproliferation in the human immune system.
Surprisingly, the lack of MHC class I expression by NOD-scid-B2mnull host cells did not appear to affect the activation or proliferation of engrafted human CD8+ cells. These data suggest that direct Ag presentation via MHC class I by mouse APC to human CD8+ cells is probably not the mechanism by which human CD8+ cells are activated. This conclusion is consistent with reports by others that indirect Ag presentation is the primary mechanism responsible for the proliferation of human T cells in a human-mouse xenogeneic mixed lymphocyte culture (56) and in the generation of human CTLs (57). Recently, however, the role of T-T Ag presentation has been highlighted (58, 59). Because the human T cells engrafted in scid mice are activated and express MHC class II molecules, T-T cell Ag presentation may also be important in their activation, proliferation, and development of an anergic phenotype. Alternatively, the costimulation and activation of human T cells may be dependent on murine host APC, as our previous data have shown that anti-mouse CD40 mAb also prevents engraftment of human T cells (20). These data, taken together, suggest that activation of human T cells may be dependent on indirect or T-T Ag presentation but independent of coactivation between human CD154 and human CD40.
It should be cautioned, however, that most human lymphocytes in scid mice are chronically activated, an observation noted in our present studies and in previously reported studies (4, 60). This suggests that CD8+ cell regulation in our model system may be mediated by memory/activated secondary responses rather than primary responses of naive T cells. Overall, based on the present data, we propose that the Hu-PBL-NOD-scid model may be suitable for the in vivo analysis of human CD4+ and CD8+ T cells and of immune regulation of EBV-related LPD development.
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2 Address correspondence and reprint requests to Dr. Dale L. Greiner, Department of Medicine, Diabetes Division, University of Massachusetts Medical School, 373 Plantation Street, Biotech 2, Suite 218, Worcester, MA 01605.
3 Abbreviations used in this paper: LPD, lymphoproliferative disorders; NOD-scid-B2mnull, NOD-scid ß2-microglobulin-null; B-LCL, B-lymphoblastoid cells; CFSE, 5-(and-6)-carboxyfluorescein succinimidyl esters; EBER, EBV-related early antigen RNA; MST, median survival time.
Received for publication December 1, 1999.
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) or in
the presence of plate-bound anti-human CD3 plus anti-human CD28
mAb (
), recombinant human IL-2 (
), or both plate-bound
anti-human CD3 plus anti-human CD28 mAb and IL-2 (
). The
cells were harvested on day 6 of culture following incubation for the
last 16 h in the presence of 1 µCi of
[3H]thymidine. Values are the mean ± SEM. *,
Significantly different from unstimulated and Ab-stimulated cultures,
p < 0.005.
