Patent Application
20200095321
Allogeneic hematopoietic cell transplantation (HCT) is a curative therapy for hematological malignancies (i.e. leukemia and lymphoma), owing to graft versus leukemia/lymphoma (GVL) effects mediated by alloreactive T cells. These same T cells also mediate acute graft-versus-host disease (GVHD) and the subsequent development of chronic GVHD (1-9). Both alloreactive CD4+ and CD8+ T cells can mediate acute GVHD, and Th1 and Th17 cells play a critical role in initiating gut GVHD (10-14). While flow cytometry-sorted donor CD4+ T cells mediate severe GVHD through expression of FASL and production of proinflammatory cytokines (i.e. IFN-γ and TNF-α) (14-17), sorted donor CD8+ T cells prevent graft rejection and mediate GVL effects through their expression of perforin/granzyme, without causing acute clinical GVHD in several mouse models (2, 18, 19). However, the mechanisms whereby purified alloreactive CD8+ T cells mediate GVL effect without causing GVHD remains largely unknown.
Programmed death ligand 1(PD-L1, also known as B7H1) functions as an immune checkpoint that interacts with PD-1 and CD80 (20, 21). PD-L1 is usually expressed by hematopoietic cells and by parenchymal cells under inflammatory cytokine (i.e. IFN-γ) induction (22). CD80 is constitutively expressed by T cells and is upregulated early after T cell activation (23), whereas PD-1 is expressed by T cells late after T cell activation (24). PD-L1 interaction with PD-1 induces anergy, exhaustion and apoptosis of activated T cells (25, 26); on the other hand, PD-L1/CD80 interaction has been reported to inhibit CD28/CTLA4 deficient T cell proliferation in vitro (21).
Expression of PD-L1 in recipient tissues decreases the severity of GVHD in conventional TBI-conditioned allogeneic recipients (27-29), while expression of PD-L1 by donor T cells increases the severity of GVHD by augmenting the expansion and survival of donor CD4+ and CD8+ T cells (30). It was shown that the interaction of PD-L1 with CD80 in the absence of PD-1 worsened GVHD by augmenting alloreactive CD4+ T cell proliferation and expansion, although simultaneous interactions of PD-L1 with both CD80 and PD-1 ameliorated GVHD by augmenting apoptosis of activated alloreactive CD4+ T cells (31).
Regulation of anergy, exhaustion, and apoptosis through PD-L1 interactions with CD80 and PD-1 on CD8+ T cells in allogeneic HCT has not yet been well characterized. It was shown that the absence of host tissue expression of PD-L1 contributed to expansion of infiltrating CD8+ T cells in GVHD target tissues in recipients with GVHD and lymphopenia (27). Other publications have shown that host tissue expression of PD-L1 caused exhaustion of alloreactive CD8+ T cells and reduced GVL effects in GVHD recipients (32, 33). However, it was reported that in vivo expansion of alloreactive CD8+ T cells in lymphoid tissues (i.e., spleen) early after HCT, before the onset of GVHD, was not affected by host tissue expression of PD-L1 (34).
The role of IFN-γ in acute GVHD pathogenesis remains controversial. IFN-γ is required for CD430 T-mediated acute GVHD in the gut and liver by augmenting Th1 differentiation and up-regulating Th1 expression of gut and liver-homing chemokine receptors (α4β7, CCR9, CCR5 and CXCR3) (29, 43, 44). In contrast, as compared to CD430 T cells, the same number of donor CD8+ Tc1 cells induced little gut acute GVHD (11, 64). IFN-γ-produced by CD8+ T cells is required to separate GVHD from GVL effects mediated by the CD8+ T cells, although IFN-γ does not directly kill tumor cells (65, 66).
IFN-γ is the key cytokine regulates tissue expression of programmed death-ligand 1 (PD-L1, also known as B7H1) (22, 67). Under non-inflammatory conditions, hematopoietic cells and lymphocytes constitutively express PD-L1 mRNA and protein, while parenchymal cells express PD-L1 mRNA without protein expression (22). Proinflammatory cytokines such as IFN-γ augment expression of PD-L1 mRNA and protein by hematopoietic cells, lymphocytes and parenchymal cells (22). Receptors for PD-L1 include CD80 and PD-1 (20, 21). PD-L1 interaction with its receptors PD-1 and CD80 induces anergy, exhaustion and apoptosis in activated T cells (25, 26). Previous studies have shown that recipient tissue expression of PD-L1 down-regulates GVHD in conventional TBI-conditioned allogeneic HCT, although the recipients still developed GVHD (29, 27, 28). It has been reported that interaction of PD-L1 with CD80 in the absence of PD-1 augmented acute GVHD by increasing alloreactive CD430 T cell proliferation without increasing CD430 T cell apoptosis, whereas simultaneous interactions of PD-L1 with both CD80 and PD-1 ameliorated GVHD by augmenting alloreactive CD430 T cell proliferation and apoptosis (31).
Accordingly, there remain needs to improve in vivo expansion of CD8+ T cells and to prevent and treat not only acute GVHD but also chronic GVHD. This invention satisfies the needs in the art.
In one aspect, the disclosure provided herein relates to a method of augmenting expansion of donor CD8+ T cells in vivo after hematopoietic cell transplantation (HCT). The method entails administering one or more doses of an effective amount of a therapeutic agent to a recipient immediately before, during, or immediately after HCT to temporarily deplete CD430 T cells or to temporarily reduce serum IL-2. In some embodiments, the therapeutic agent includes an anti-CD4 antibody or an anti-CD4-meditope-immunotoxin. In some embodiments, the anti-CD4+ antibody is a monoclonal antibody or a humanized antibody. In some embodiments, the therapeutic agent includes an anti-IL-2 antibody (e.g., an anti-IL-2 monoclonal antibody and/or humanized antibody) or an agent blocking IL-2R. In some embodiments, the CD8+ T cells are selectively expanded in lymphoid tissues but not in GVHD target tissues of the subject. In some embodiments, the expanded CD8+ T cells produce an increased amount of IFN-, comparing to control recipients received with IgG.
In another aspect, the disclosure provided herein relates to a method of preventing a subject from suffering from GVHD or treating a subject suffering from GVHD after HCT while preserving GVL. The method entails administering one or more doses of an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT to temporarily deplete CD430 T cells or to temporarily reducing serum IL-2. In some embodiments, the therapeutic agent includes, but is not limited to, an anti-CD4 antibody, an anti-CD4-meditope-immunotoxin, an anti-IL-2 antibody, or an IL-2R blocking agent. In some embodiments, the anti-CD4+ antibody is a monoclonal antibody or a humanized antibody. In some embodiments, acute GVHD is prevented or treated by administering to the subject a single dose of the therapeutic agent. In some embodiments, GVHD is prevented or treated by administering no more than three doses of the therapeutic agent. For example, the three doses are administered within one month, at one- or two-week intervals. In some embodiments, more than three doses of the therapeutic agent can be administered to prevent or treat GVHD. In some embodiments, one or more doses of PD-L1-Ig are administered to prevent or treat GVHD while preserving GVL. In some embodiments, the method further entails administration of one or more doses of IFN- to the subject in addition to temporarily depleting CD430 T cells or reducing serum IL-2.
In another aspect, the disclosure provided herein relates to a method of preventing or treating GVHD and augmenting thymus recovery after HCT. The method entails administering one or more doses of an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT to temporarily deplete CD430 T cells from the transplant and from de novo generation or to temporarily reduce serum IL-2 for a period from 60 days to 120 days. In some embodiments, the therapeutic agent includes an anti-CD4 antibody, or an anti-CD4-meditope-immunotoxin. In some embodiments, the anti-CD4+ antibody is a monoclonal antibody or a humanized antibody. In some embodiments, the therapeutic agent includes an anti-IL2 antibody, or an agent blocking IL-2R. In some embodiments, the anti-IL2 antibody is a monoclonal antibody or a humanized antibody.
In another aspect, the disclosure provided herein relates to a method of augmenting recipient tissue expression of programmed death-ligand 1 (PD-L1, or B7H1) after HCT. The method entails administering one or more doses of an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT. In some embodiments, the therapeutic agent includes an agent that temporarily depletes CD430 T cells, such as an anti-CD4 antibody (e.g., a monoclonal or humanized anti-CD4 antibody) or an anti-CD4-meditope-immunotoxin. In some embodiments, the therapeutic agent includes an agent that temporarily reduces serum IL-2, such as an anti-IL-2 antibody (e.g., a monoclonal or humanized anti-IL-2 antibody) or an agent blocking IL-2R.
This application contains at least one drawing executed in color. Copies of this application with color drawing(s) will be provided by the Office upon request and payment of the necessary fees.
concentrations but decreased IL-2 concentrations and augmented CD8+ T cell expansion in lymphoid tissues but not in GVHD target tissues. BALB/c recipients transplanted with splenocytes (2.5×106) and TCD-BM cells from C57BL/6 donors were injected with either rat IgG or anti-CD4 mAb (500 μg/mouse) at day 0 after HCT.
−/− donor transplants. Lethally irradiated BALB/c recipients transplanted with splenocytes (5×106) and TCD-BM (2.5×106) from wild-type or IFN-
−/− C57BL/6 donors, and then given a single i.v. injection of anti-CD4 mAb (500 μg/mouse) at the time of HCT. Recipients were monitored for clinical signs of GVHD, including body weight change, diarrhea, hair loss, and survival.
The following description of the invention is merely intended to illustrate various embodiments of the invention. As such, the specific modifications discussed are not to be construed as limitations on the scope of the invention. It will be apparent to one skilled in the art that various equivalents, changes, and modifications may be made without departing from the scope of the invention, and it is understood that such equivalent embodiments are to be included herein.
Many factors and pathways may contribute to acute GVHD or chronic GVHD after HCT. Disclosed herein are methods of preventing or treating acute GVHD and chronic GVHD while preserving GVL effects, methods of augmenting expansion of donor CD8+ T cells in lymphoid tissues in vivo after HCT, and methods of augmenting recipient tissue expression of PD-L1 after HCT. GVHD prevention and treatment, as well as in vivo expanding donor CD8+ T cells, can be achieved by temporarily depleting CD430 T cells using an anti-CD4 agent such as an anti-CD4 antibody or an anti-CD4-meditope-immunotoxin, neutralizing IL-2 using an anti-IL2 antibody, or administering other agents blocking IL-2R. Moreover, other therapeutic agents, such as PD-L1 antibodies and/or IFN- can be administered to the subject receiving HCT.
PD-L1 interacts with PD-1 and CD80, and functions as a checkpoint that regulates immune responses in animal models and humans. It is disclosed herein that in allogeneic and xenogeneic murine models of graft-versus-host disease (GVHD), temporary depletion of donor CD4+ T cells immediately after hematopoietic cell transplantation (HCT) effectively prevents GVHD while preserving strong graft-versus-leukemia (GVL) effects. Depletion of donor CD4+ T cells increases serum IFN-γ but reduces IL-2 concentrations, leading to upregulated expression of PD-L1 by recipient GVHD target tissues and by donor CD8+ T cells. In GVHD target tissues, the interactions of PD-L1 with PD-1 on donor CD8+ T cells induced tolerance through anergy, exhaustion and apoptosis of effector T cells, thereby preventing GVHD. In lymphoid tissues, the interactions of PD-L1 with CD80 augment CD8+ T cell expansion and activity against malignant cells in the recipient, without increasing anergy, exhaustion or apoptosis, resulting in strong GVL effects. These results show that the outcome of PD-L1-mediated signaling in CD8+ T cells depends on the presence or absence of CD4+ T cells, the nature of the interacting receptor expressed by CD8+ T cells, and the tissue environment where the signaling occurs.
As detailed in this disclosure, augmenting CD8+ T cells in lymphoid tissues as well as expressing PD-L1 in recipient tissues by administering a therapeutic agent to the recipient has unexpectedly prevented or treated not only acute GVHD but also chronic GVHD. Surprisingly, a single dose of the therapeutic agent is sufficient to prevent or treat acute GVHD and as few as three doses of the therapeutic agent administered within one month are sufficient to prevent or treat chronic GVHD.
The term “recipient,” “host,” “subject,” or “patient” as used herein refers to a subject receiving hematopoietic cell transplantation. These terms may refer to, for example, a subject receiving an administration of donor bone marrow, donor T cells, donor spleen cells, or other donor cells or tissue. In some embodiments, the transplanted cells are derived from an allogeneic donor. The recipient, host, subject, or patient can be an animal, a mammal, or a human.
The term “donor” as used herein refers to a subject from whom the cells or tissue are obtained to be transplanted into a recipient or host. For example, a donor may be a subject from whom bone marrow, T cells, spleen cells, or other cells or tissue to be administered to a recipient or host is derived. The donor or subject can be an animal, a mammal, or a human.
The terms “treat,” “treating,” and “treatment” as used herein with regard to a GVHD condition refer to alleviating the condition partially or entirely, or eliminating, reducing, or slowing the development of one or more symptoms associated with the condition. In some embodiments, the term “treat,” “treating,” or “treatment” means that one or more symptoms of GVHD condition or complications are alleviated in a subject receiving the treatment as disclosed herein comparing to a subject who does not receive such treatment.
The terms “prevent,” “preventing,” and “prevention” as used herein with regard to a GVHD condition refer to preventing the onset of the condition and/or symptoms associated with the condition from occurring, decreasing the likelihood of occurrence or recurrence of the condition, or slowing the progression or development of the condition.
The phrase “an effective amount” or “a therapeutically effective amount” as used herein refers to an amount of a therapeutic agent that produces a desired therapeutic effect. For example, an effective amount of an anti-CD4 antibody may refer to that amount that prevents or treats GVHD, depletes CD430 T cells, augments CD8+ T cells, or induces tissue expression of PD-L1 in a recipient. The precise effective amount is an amount of the therapeutic agent that will yield the most effective results in terms of efficacy in a given subject. This amount will vary depending upon a variety of factors, including but not limited to the characteristics of the therapeutic agent (including activity, pharmacokinetics, pharmacodynamics, and bioavailability), the physiological condition of the subject (including age, sex, disease type and stage, general physical condition, responsiveness to a given dosage, and type of medication), the nature of the pharmaceutically acceptable carrier or carriers in the formulation, and the route of administration. One skilled in the clinical and pharmacological arts will be able to determine a therapeutically effective amount through routine experimentation, namely by monitoring a subject's response to administration of a compound and adjusting the dosage accordingly. For additional guidance, see Remington: The Science and Practice of Pharmacy (Gennaro ed. 20th edition, Williams & Wilkins PA, USA) (2000).
In one aspect, the disclosure provided herein relates to a method of preventing or treating chronic GVHD after HCT while preserving GVL. The method entails in vivo administering two or more doses of an effective amount of a therapeutic agent to a recipient simultaneously or immediately after HCT to temporarily deplete CD430 T cells.
The term “simultaneously” as used herein with regards to administration means that the therapeutic agent is administered to the recipient at the same time or nearly at the same time of HCT. For example, the therapeutic agent is considered to be administered “simultaneously” if it is administered via a single combined administration of hematopoietic cells, two or more administrations occurring at the same time, or two or more administrations occurring in succession without extended intervals in between.
When the therapeutic agent is administered immediately before HCT, a first dose of the therapeutic agent can be administered any time up to about 10 days before HCT. When the therapeutic agent is administered immediately after HCT, a first dose of the therapeutic agent can be administered any time up to about 6 weeks after HCT. In some embodiments, a first dose of the therapeutic agent is administered about 1 hour, about 2 hours, about 3 hours, about 4 hours, about 5 hours, about 6 hours, about 7 hours, about 8 hours, about 9 hours, about 10 hours, about 11 hours, about 12 hours, about 24 hours, about 2 days, about 3 days, about 4 days, about 5 days, about 6 days, about 7 days, about 8 days, about 9 days, or about 10 days, before HCT. In some embodiments, a first dose of the therapeutic agent is administered simultaneously with HCT. In some embodiments, a first dose of the therapeutic agent is administered about 1 hour, about 2 hours, about 3 hours, about 4 hours, about 5 hours, about 6 hours, about 7 hours, about 8 hours, about 9 hours, about 10 hours, about 11 hours, about 12 hours, about 24 hours, about 2 days, about 3 days, about 4 days, about 5 days, about 6 days, about 7 days, about 8 days, about 9 days, about 10 days, about 11 days, about 12 days, about 13 days, about 14 days, about 3 weeks, about 4 weeks, about 5 weeks, or about 6 weeks, after HCT.
When multiple doses of the therapeutic agent are administered, it is within the purview of one of ordinary skill in the art to adjust the administration schedule to optimize the therapeutic effect. For example, one dose can be administered immediately before HCT, followed by additional doses administered during and/or immediately after HCT. In some embodiments, one or more doses of the therapeutic agent can be administered subsequently after the administration of the first dose, e.g., within one month of administration of the first dose. For example, the subsequent doses of the therapeutic agent can be administered in one-week intervals or in two-week intervals.
By administering one or more doses of the therapeutic agent depleting CD4+ T cells immediately after HCT, the donor CD430 T cells as well as de novo generated CD430 T cells are completely and temporarily depleted. For example, at least 90%, at least 95%, at least 98%, or at least 99% of the CD430 T cells are depleted. The CD4+ T cells are depleted for only a short period of time, for less than 10 weeks, for less than 9 weeks, for less than 8 weeks, for less than 7 weeks, for less than 6 weeks, for less than 5 weeks, for less than 4 weeks, for less than 3 weeks, or for about two weeks. In some embodiments, the CD430 T cells are depleted for at least two weeks.
Any therapeutic agent that effectively depletes CD430 T cells in vivo for a temporary period of time can be used. In some embodiments, the therapeutic agent is an anti-CD4 antibody, preferably a monoclonal antibody or a humanized antibody. For example, a depleting anti-human CD4 mAb is disclosed in U.S. Pat. No. 8,399,621, the content of which is incorporated herein by reference in its entirety. A functional fragment of an anti-CD4 antibody can be used as long as the fragment effectively depletes CD430 T cells in vivo. In some embodiments, CD430 T cells can be depleted by administering to the subject an anti-CD4-meditope-immunotoxin. Such meditopes can be made according to technology known in the art (68). It is within the purview of one of ordinary skill in the art to determine the dose of the therapeutic agent to achieve a desired duration period of depleting CD430 T cells in vivo.
In some embodiments, a therapeutic agent that effectively neutralizes IL-2 in vivo for a temporary period of time can be used. Such agents include but are not limited to anti-IL-2 antibody, including monoclonal antibodies and/or humanized antibodies, or other agents blocking IL-2R. Certain anti-IL-2 receptor antibodies are known in the art (76, 77). It was reported that IL-2 administration was able to prevent acute GVHD or chronic GVHD (69, 70). Surprisingly, administration of an IL-2 antibody is effective in preventing or treating acute GVHD, as disclosed herein.
In some embodiments, a therapeutic agent includes a PD-L1-Ig. Administration of one or more doses of a therapeutically effective amount of a PD-L1-Ig can also prevent or treat GVHD.
In some embodiments, one or more doses of IFN- can be administered to the subject in the absence of CD430 T cells or at a reduced serum level of IL-2 to help preserve GVL.
In another aspect, the disclosure provided herein relates to a method of preventing or treating acute GVHD after HCT while preserving GVL. The method entails in vivo administering an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT to temporarily deplete CD430 T cells or to temporarily reduce the serum IL-2. Surprisingly, only a single dose of the therapeutic agent is sufficient to prevent or treat acute GVHD. In some embodiments, a single dose of an anti-CD4 antibody is sufficient to prevent or treat acute GVHD. In some embodiments, multiple doses of an anti-IL-2 antibody is administered. For example, an anti-IL-2 antibody can be injected to a subject receiving HCT every other day for up to 30 days to effectively prevent gut GVHD.
In some embodiments, the single dose of the therapeutic agent is administered to the recipient simultaneously with HCT, as described above. Or the single dose of the therapeutical agent is administered immediately before or immediately after HCT, as described above.
In another aspect, the disclosure provided herein relates to a method of augmenting expansion of donor CD8+ T cells in lymphoid tissues in vivo after HCT. The method entails in vivo administering an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT to temporarily deplete CD430 T cells or to temporarily reduce serum IL-2.
In recipient lymphoid tissues, donor CD8+ T cell proliferation is augmented without increasing CD8+ T cell anergy or apoptosis, thereby to achieve strong GVL effects. Surprisingly, in GVHD target tissues, anergy and apoptosis of infiltrating CD8+ T cells are increased in a manner dependent on recipient PD-L1 expression, thereby preventing damage to intestinal Paneth cells and stem cells, hepatocytes, and thymic medullary epithelial cells.
In another aspect, the disclosure provided herein relates to a method of augmenting recipient tissue expression of programmed death-ligand 1 (PD-L1, or B7H1) after HCT. The method entails administering an effective amount of a therapeutic agent to a recipient simultaneously, immediately before, or immediately after HCT to temporarily deplete CD430 T cells or to temporarily reduce serum IL-2.
Although down-regulation of GVHD by host tissue PD-L1 has been previously reported with the use of PD-L1 deficient recipients (25, 26), this disclosure for the first time demonstrates that host tissue expression of PD-L1 can be achieved by administration of a CD430 T cells-depleting agent immediately after HCT to effectively prevent both acute and chronic GVHD and preserve strong GVL effects. Depletion of donor CD430 T cells increases serum IFN-γ concentration and enhances recipient tissue expression of PD-L1 and donor CD8+ T cell expression of the PD-L1 receptors CD80 and PD-1.
As detailed in this disclosure, temporary in vivo depletion of donor CD430 T cells immediately after transplantation increases donor CD8+ T cell susceptibility to anergy and apoptosis, an effect mediated by expression of PD-L1 in GVHD target tissues. Host expression of PD-L1 has little effect on donor CD8+ T cell expansion in the lymphoid tissues immediately after HCT. Temporary in vivo depletion of donor CD430 T cells augments donor CD8+ T expansion and allows potent CD8+ T cell-mediated GVL effects in recipient lymphoid tissues. PD-L1 interactions with CD80 and PD-1 mediates donor CD8+ T cell anergy, exhaustion, and apoptosis in different GVHD target tissues in a tissue-specific manner.
With murine models of allogeneic GVHD that reflects characteristic features of acute and chronic GVHD in humans and a murine model of xenogeneic GVHD induced by human PBMC (11, 39, 53), the working examples disclosed herein demonstrate that temporary in vivo depletion of adoptively transferred mature donor CD430 T and de novo-generated CD430 T cells immediately after HCT prevents both acute and chronic GVHD while augmenting donor early CD8+ T expansion in lymphoid tissues and preserving strong GVL effects. This outcome does not simply reflect depletion of donor CD4+ T cells that recognize recipient alloantigens, but results from several newly observed mechanisms. Depletion of donor CD4+ T cells leads to increase of serum IFN- and decrease of IL-2 concentrations. Depletion of donor CD4+ T cells also leads to expansion of donor CD8+ T cells via T-T and PD-L1/CD80 interactions in lymphoid tissues where they mediate strong GVL effects. At the same time, depletion of donor CD4+ T cells enables host-tissue expression of PD-L1 to induce anergy, exhaustion, and apoptosis of CD8+ T cells infiltrating GVHD target tissues via PD-L1/PD-1 interactions in a tissue-specific manner.
Expression of PD-L1 in recipient tissues can prevent both acute and chronic GVHD after effective depletion of donor CD430 T cells immediately after HCT, and temporary depletion for only 30-60 days after HCT is sufficient. As demonstrated in the working examples, a single injection of anti-CD4 effectively prevented acute GVHD, but the recipients still developed chronic GVHD with damage in GVHD target tissues, especially in the salivary gland. The working examples further demonstrate that at least three injections were required to effectively prevent chronic GVHD. Three injections of anti-CD4 allowed medullar thymic epithelial cell (mTEC) recovery and restoration of thymic negative selection, but a single injection was not sufficient. It was reported that de novo generated CD430 T cells immediately after HCT could perpetuate CD8+-mediated damage in the thymus, leading to autoimmune-like chronic GVHD (11). Although a single injection of anti-CD4 prevented acute GVHD and augmented de novo generation of donor-type CD430 T cells, it did not prevent thymus damage mediated by de novo-generated donor CD430 T cells immediately after HCT. On the other hand, in the absence of donor CD430 T cells, donor CD8+ T cells infiltrating thymic tissues were tolerized by host-tissue PD-L1, and thymus damage-mediated by the donor CD8+ T cells was self-limited. Therefore, anti-CD4 treatment has the important effect of temporarily depleting both the injected mature CD430 T cells and also the CD430 T cells generated de novo from the marrow early after HCT, thereby allowing sufficient time for mTEC to recover and restore effective thymic negative selection. This time period is proximately 30-60 days after HCT. CD430 T cells generated from the donor marrow after this time point no longer cause chronic GVHD.
Clinical GVHD prevention is usually associated with reduction of alloreactive T cell expansion and proinflammatory cytokine (i.e. IFN- and TNF-α) production. The working examples demonstrate that a single injection of depleting anti-CD4 immediately after HCT effectively prevented acute GVHD, even though the depletion of donor CD4+ T cells led to strikingly increased serum IFN-
concentrations immediately after transplantation. These results were unexpected since IFN-
contributes to the pathogenesis of gut GVHD and exacerbates GVHD after PD-1 blockade in recipients transplanted with both donor CD4+ and CD8+ T cells (41, 54). On the other hand, these results are consistent with results reported by Yang et al. (55) who showed that in the absence of donor CD4+ T cells, IFN-γ-deficient donor CD8+ T cells proliferated more vigorously and caused more severe GVHD than WT donor CD8+ T cells.
As shown in the working examples, increased IFN- concentrations were associated with enhanced expression of PD-L1 by colon epithelial cells and IFN-
deficient donor cells was associated with down-regulation of PD-L1 expression. These observations are consistent with results of a previous study showing that in recipients with acute GVHD, upregulation of PD-L1 expression in host tissues requires IFN-
(29). Although host tissue PD-L1 had little impact on donor CD8+ T cell proliferation or survival in the spleen immediately after HCT, depletion of donor CD4+ T cells led to induction of apoptosis of infiltrating donor CD8+ T cells by PD-L1 in colon tissue and induction of anergy and exhaustion by PD-L1 in liver tissue. The differential effect of PD-L1-mediated signaling on donor CD8+ T cells in the colon and liver was associated with differential expression of PD-1 and CD80 by donor CD8+ T cells in these tissues. The ratio of PD-1 versus CD80 MFI on donor CD8+ T cells was significantly higher in the colon as compared to the liver.
NKT cells, myeloid suppressor cells (MDSCs), and regulatory T cells can suppress GVHD (5, 58) and some of these cells express CD4 and could be depleted by anti-CD4-treatment. However, the working examples demonstrate that depletion of donor CD430 T cells together with those CD4+ regulatory cells was able to effectively prevent GVHD, suggesting that in the absence of donor CD430 T cells, tissue protective mechanisms are sufficient to prevent GVHD mediated by CD8+ T cells, and CD4+ regulatory T cells are dispensable.
The working examples demonstrate that small numbers of donor CD430 T cells in the graft could augment acute GVHD by markedly reducing the apoptosis of CD8+ T cells infiltrating the colon, and this effect was IL-2 dependent. In addition, although sorted CD8+ T cells induced little GVHD, and sorted CD430 T cells induced severe acute GVHD in PD-L1 sufficient wild-type recipients, sorted CD8+ and CD430 T cells both induced lethal acute GVHD with similar severity in PD-L1-deficient recipients. These results suggest that CD8+ T cells are more sensitive than CD430 T cells to host-tissue PD-L1-mediated apoptosis, and CD430 T cell help immediately after HCT can make donor CD8+ T cells resistant to host-tissue PD-L1-mediated apoptosis. This observation is consistent with a previous report that IL-2 from CD4+ T cells may prevent apoptosis induced by PD-1 signaling in CD8+ T cells that are deficient in IL-2 production (59). These results support that host-tissue expression of PD-L1 could ameliorate GVHD only to a certain degree when whole spleen cells with both CD4+ and CD8+ T cells were transplanted, as indicated by comparing WT and PD-L1−/− recipients (27, 28). As demonstrated in the working examples, increased host tissue expression of PD-L1 induced by higher concentrations of IFN-γ combined with increased sensitivity of donor CD8+ T cells to PD-L1-induced apoptosis in the presence of lower IL-2 concentrations could explain the highly effective prevention of GVHD that was found when donor CD4+ T cells were depleted immediately after transplantation. Depletion of donor CD4+ T cells immediately after HCT not only prevented GVHD, but also enabled donor CD8+ T cell expression of PD-L1 to mediate their own expansion in lymphoid tissues and mediate strong GVL activity that could overcome “GVL-resistant” BC-CML tumor cells. The high concentrations of IFN-γ associated with CD4+ T cell depletion could contribute to the preservation of GVL activity, since Yang et al. (55) showed that in the absence of donor CD430 T cells, IFN-γ-deficient donor CD8+ T cells had lower GVL activity than WT donor CD8+ T cells.
The working examples demonstrate that anti-CD4-treatment immediately following HCT augments donor CD8+ T cell expansion in the lymphoid tissues, which is dependent on donor CD8+ T expression of both PD-L1 and CD80, and host-tissue expression of PD-L1 has little impact. The lack of impact from host PD-L1 is likely due to relative paucity of host parenchymal cells that express PD-L1 in the lymphoid tissues. The expansion of donor CD8+ T cells in lymphoid tissues most likely results from T-T interaction via PD-L1/CD80, although the possibility that CD8+ T interaction with non-T cells via PD-L1/CD80 cannot be excluded. First, PD-L1 deficiency on donor CD8+ T cells, but not PD-L1 deficiency on donor non-T cells (data not shown) markedly reduced donor CD8+ T cell survival and expansion. Second, CD80 deficiency on donor CD8+ T cells also reduced donor CD8+ T expression of survival gene BCL-XL and increased CD8+ T cell exhaustion. Third, anti-CD4-treatment immediately after HCT upregulated PD-L1 and CD80 expression by donor CD8+ T cells but not by non-T cells (i.e. DCs and myeloid cells). Finally, specific blockade of PD-L1/CD80 interaction by anti-PD-L1 (43H12) markedly reduced donor CD8+ T survival and expansion in the spleen and abolished GVL effects. This result is consistent with previous findings that PD-L1 on CD8+ T cells was required for survival of activated CD8+ T cells (60). In contrast, PD-L1/CD80 interactions augment apoptosis of activated CD4+ T cells (31).
Saha et al showed that PD-L1 deficiency in donor T cells reduced proliferation and survival of donor T cells and delayed GVHD lethality in recipients given both CD4+ and CD8+ donor T cells (30). This observation suggests that even when donor T cells have reduced survival capacity due to lack of expression of PD-L1, host tissue expression of PD-L1 is still unable to tolerize tissue infiltrating T cells to prevent GVHD when both donor CD4+ and CD8+ T cells were present. The working examples show that depletion of donor CD4+ T cells immediately after HCT allowed host PD-L1 to effectively tolerize infiltrating CD8+ T cells in GVHD target tissues and completely prevent acute GVHD. At the same time, PD-L1/CD80 interactions among donor CD8+ T cells in the lymphoid tissues augment CD8+ T cell survival and expansion as well as their GVL activity.
On the other hand, using a non-lethal murine model of GVHD mediated by H-Y antigen-specific transgenic CD8+ T cells, Michonneau et al reported that the transgenic CD8+ T cells were not able to eliminate host-type tumor cells in lymphoid tissues due to enhanced PD-L1/PD-1 interactions between PD-1+ transgenic CD8+ T cells and PD-L1+ CD11c+ DCs and F4/80+ macrophages (52). Similarly, Mueller et al showed that PD-L1 expressed by hematopoietic cells suppressed viral-specific CD8+ cell activation and expansion (61). Although the working examples show that CD11+ DCs and MAC-1/Gr-1+ myeloid cells in the spleen expressed much higher levels of PD-L1 as compared to those in the liver, the high levels of PD-L1 on DCs and myeloid cells did not appear to interfere with the GVL activity of donor CD8+ T cells in the lymphoid tissues of anti-CD4-treated recipients, since different types and dose of tumor cells were all eliminated in anti-CD4-treated recipients.
Several explanations might account for the different impact of PD-L1-expressed by hematopoietic cells on GVL effects in the lymphoid tissues of recipients given H-Y-specific CD8+ T cells as compared to wild-type alloreactive CD8+ T cells. First, PD-L1 expressed by hematopoietic cells mainly control activation and expansion of naïve T cells (61). In anti-CD4-treated recipients, alloreactive CD8+ T cells are activated by recipient APCs that are rapidly eliminated. Therefore, PD-L1 expression by donor hematopoietic-derived APCs does not play an important role on donor T cell activation and expansion. Second, H-Y-specific transgenic CD8+ T cells in male recipients appeared to have very weak alloreactivity as indicated by lack of GVHD mortality even after blockade of PD-1. Their alloreactivity was easily controlled by PD-L1/PD-1 interactions between CD8+ T cells and DCs and macrophages in the lymphoid tissues. In contrast, the alloreactivity of wild-type alloreactive CD8+ T cells is much stronger, as indicated by their ability to cause rapidly lethal GVHD in PD-L1−/− recipients. Their alloreactivity cannot be controlled by PD-L1/PD-1 interactions between CD8+ T and DCs and macrophage. Third, H-Y-specific transgenic CD8+ T cells might not express PD-L1, or PD-L1 might not play a role in their survival and expansion, unlike wild-type alloreactive T cells (30).
As depicted in but decreases IL-2 concentrations. Increase of IFN-
augments expression of PD-L1 by donor CD8+ T cells and host tissues, while increasing expression of PD-1 and CD80 by donor CD8+ T cells. Donor CD8+ T cells express higher levels of PD-L1 and CD80 but lower level of PD-1 in the spleen, promoting PD-L1/CD80 interactions among donor CD8+ T cells. In contrast, donor CD8+ T cells express higher level of PD-1 and lower levels of PD-L1 and CD80 in GVHD target tissues, promoting host tissue PD-L1 interaction with PD-1 on donor CD8+ T cells. CD8+ T cells are defective in IL-2 production, and in the absence of IL-2 help from CD4+ T cells, donor CD8+ T cells may become more sensitive to the tolerizing effects of PD-L1/PD-1 signaling. Donor CD8+ T-T and PD-L1/CD80 interactions augment donor CD8+ T survival and expansion in lymphoid tissues, resulting in strong GVL effects. Dominant host-PD-L1 interaction with PD-1 on CD8+ T cells mediates donor CD8+ T cell anergy, exhaustion and apoptosis in GVHD target tissues, thereby preventing GVHD.
The working examples support that sorted donor CD8+ T cells facilitate engraftment and mediate GVL effect without causing GVHD (2, 7). The results also demonstrate that ex vivo depletion of donor CD430 T cells did not effectively prevent GVHD in a previous human trial (62) probably because very small numbers of donor CD430 T cells in the graft could have expanded after HCT, and they could have worked together with donor CD430 T cells generated from the marrow progenitors immediately after HCT to help donor CD8+ T cells resist host-tissue PD-L1 mediated apoptosis or other tolerance mechanisms. The results also indicate that temporary in vivo depletion of donor CD430 T cells immediately after HCT can be a novel approach to prevent GVHD while preserving strong GVL effects. Temporary in vivo depletion of donor CD430 T cells for a period of approximately 30-60 days after HCT may not only allow GVHD target tissues to tolerize infiltrating donor CD8+ T cells while preserving GVL effects in the lymphoid tissues, but may also allow regeneration of medullary thymic epithelia cells and restoration of thymic negative selection for durable prevention of chronic GVHD.
Permanent depletion of de novo-generated CD430 T cells is not required for mTEC recovery. The results in the working examples indicate that recovery of mTECs takes time. Autoreactive CD430 T cells can be generated de novo immediately after HCT before mTEC have adequately recovered, but as time goes on, the mTEC percentage gradually increases, and negative selection is gradually restored. Based on the results disclosed herein, CD430 T cells generated de novo beyond ˜45 days after HCT no longer cause autoimmunity or chronic GVHD. Therefore, depletion of de novo-generated autoreactive CD430 T cells immediately after HCT allows time for mTEC recovery and restoration of negative selection in the thymus. The methods disclosed herein should not cause long-term CD430 T cell deficiency in young recipients with adequate thymic function, although CD430 T cell reconstitution may be delayed in older recipients.
The following examples are provided to better illustrate the claimed invention and are not to be interpreted as limiting the scope of the invention. To the extent that specific materials are mentioned, it is merely for purposes of illustration and is not intended to limit the invention. One skilled in the art may develop equivalent means or reactants without the exercise of inventive capacity and without departing from the scope of the invention.
Induction and scoring of acute GVHD and chronic GVHD, in vivo bioluminescent imaging, in vivo BrdU-labeling of proliferating T cells, TUNEL staining, tissue cell isolation, intracellular staining of cytokines, antibodies, flow cytometry analysis and sorting, histopathology, histoimmunofluorescent staining, and real-time PCR have been described in previous publications (11, 27, 29, 31, 63) and detailed below.
Data are displayed as mean±SEM. Bodyweight, diarrhea, cutaneous damage scoring, GVHD and survival in different groups were compared by using the rank sum test or log-rank test. Comparison of two means was analyzed using an unpaired two-tailed Student t-test (Prism, version 6.0; GraphPad Software) (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0005).
Mice: C57BL/6 (H-2b) and BALB/c (H-2d) mice were purchased from the National Cancer Institute animal production program (Frederick, Md.). A/J mice (H-2a) were purchased from the Jackson Laboratory (JAX). PD-L1−/− BALB/c breeders were provided by Dr. Lieping Chen (Yale University). PD-L1−/− C57BL/6 breeders, spleen and bone marrow cells were provided by Dr. Haidong Dong (Mayo Clinic). Congenic CD45.1+ C57BL/6 mice, CD80−/− C57BL/6 breeders and IFN-γ−/− C57BL/6 breeders were purchased from JAX Lab. Rag2−/− BALB/c mice were purchased from Taconic Farms (Germantown, N.Y.). NSG mice were provided by the Animal Tumor Model Core (City of Hope). All mice were maintained in a pathogen-free room in the City of Hope Animal Resource Center. All animal protocols were approved by COH Institutional Animal care and use committee (IACUC).
Induction and assessment of GVHD: BALB/c recipients were exposed to 850 cGy total body irradiation (TBI) with the use of a [137Cs] source 8-10 hours before HCT, and then injected intravenously (i.v.) with C57BL/6 donor spleen cells (2.5×106 or 5.0×106) and T cell-depleted BM (TCD-BM) (2.5×106). C57BL/6 recipients were exposed to 1100 cGy TBI and then injected i.v. with A/J donor spleen cells (10×106, 20×106 or 40×106) or CD8+ TCD spleen and BM cells (10×106). NSG recipients were injected i.p. with human PBMC (20×106) from healthy donors. For secondary transplantation, Rag2−/− BALB/c mice were exposed to 200 cGy TBI 24 h before HCT and were injected i.v with sorted CD8+ T cells (1×106) from the liver of anti-CD4 or rat-IgG-treated primary recipients together with primary recipient strain TCD-BM (5×106). T cell depletion from the bone marrow was accomplished by using biotin-conjugated anti-CD4 and anti-CD8 mAbs, and streptavidin Microbeads (Miltenyi Biotec, Germany), followed by passage through an autoMACS Pro cell sorter (Miltenyi Biotec, Germany). Enrichment of Thy1.2+ cells from spleen was accomplished by using mouse anti-CD90.2 microbeads (Miltenyi Biotec, Germany). The purity of enrichment was >98%, whereas the purity of depletion was >99%. The assessment and scoring of clinical acute signs of GVHD and clinical cutaneous GVHD has been described previously (1, 2).
Isolations of cells from GVHD target tissues: Liver samples were mashed through a 70 μm cell strainer, and MNC were isolated from the cell suspensions with Lymphocyte M. Digestion buffer [RPMI containing 5% fetal bovine, 10 mM HEPES, 10 U heparin, collagenase D (1 mg/ml), and DNase I (1000 U/ml)] was carefully injected into lung lobes, and specimens were incubated at 37° C. for 45 min. After a second cycle of digestion, lung tissue were mashed through a 70 μm cell strainer, and MNC were isolated from cell suspensions with Lymphocyte M. Colon specimens were washed in PBS, cut into 0.5 mm pieces and suspended in PBS containing 1% Bovine serum and 0.002 M EDTA, vortexed for 10 min., passed through 70 μm strainer and glass wool, and centrifuged for 5 min at 2000 rpm to isolate epithelial cells and lymphocyte.
Antibodies, FACS analysis and FACS sorting: Purified depleting anti-mouse CD4 mAb (GK1.5), blocking anti-mouse PD-L1 (10F.9G2), neutralizing anti-IL-2 (JES6-1A12), and CD8 (53-6.72) for in vivo treatment were purchased from Bio X Cell (West Lebanon, N.H.). Depleting anti-human CD4 mAb (IT1208) for in vivo treatment was provided by Dr. Ito at IDAC Theranostics. H-2Kb (AF6-88.5), α4β7 (DATK32), Ly51 (6C3) and FITC Annexin V were purchased from BD Pharmingen (San Diego, Calif.). mAbs to TCRβ (H57-597), H-2Kb (AF6-88.5), CD3(UCHT1),CD4 (RM4-5), CD8a(SK1), CD8a (53-6.7), CD45 (30-F11), CD11b(M1/70), CD11c(N418), Gr-1(RB6-8C5), B7H1 (H1M5), PD-1 (RMP1-30), CD44 (IM7), CD62L (MEL-14), EpCAM (G8.8), FASL (MFL3), IL7Rα (A7R34), TIM3 (RMT3-23), IFN-γ (XMG1.2), EOMES (Dan 11 mag) and Foxp3 (FJK-16s) were purchased from eBioscience (San Diego, Calif.). mAbs to CCR9 (Clone 242503) and IL-22R (Clone 496514) were purchased from R&D Systems (Minneapolis, Minn.). Anti-CXCR3 mAb and anti-T-bet (4610) were purchased from Biolegend (San Diego, Calif.). Polyclonal Rabbit Anti-Human Lysozyme EC 3.2.1.17 was purchased from DAKO (Carpinteria, Calif.). Anti-RNF128:FITC (GRAIL) mAb (ARP43311_T100) were purchased AVIVA SYSTEMS BIOLOGY (San Diego, Calif.). Anti-Cytokeratin mAb was purchased from Sigma-Aldrich (Louis, MO). mAb to Ulex europaeus agglutinin 1 (UEA-1) was purchased from Vector Laboratories (Burlingame, Calif.). Flow cytometry analyses were performed with a CyAn Immunocytometry system (DAKO Cytomation, Fort Collins, Colo.) and BD LSRFortessa (Franklin Lakes, N.J.), the resulting data were analyzed with FlowJo software (Tree Star, Ashland, Oreg.). T cell sorting was performed with a BD FACS Aria SORP sorter at the City of Hope FACS facility. The sorted cells were used for transplantation and real-time RT-PCR.
GVHD target tissue cell isolation: Mononuclear cells (MNCs) from lung, liver and gut were processed and collected as previously described (29). Thymic epithelial cell isolation was performed as previously described (11). In brief, the thymus was cut into small pieces and placed in RPMI 1640 media with collagenase D and DNAse I. Thymic fragments were rapidly mixed through the aperture of a 1000-ml pipette tip and incubated in a 37° C. water bath to digest the thymus and release epithelial cells from the extracellular matrix. Cell suspension was harvested every 15 min, and the process was repeated twice. The harvested cells were incubated with anti-CD45 microbeads, followed by passing through an MACS separation column (Miltenyi Biotec), the negative population containing CD45− mTEC cells were kept for the subsequent flow cytometry analysis. The gut epithelial cell isolation was performed according to a previous report (71). Briefly, colons were washed in PBS and chopped into 0.5 cm pieces. Colon tissue was incubated in 5 mM EDTA and 1 mM DTT with PBS for 30 min at 37° C. while shaking at 200 rpm. Samples were filtered in a 70-μm strainer, centrifuged for 15 min at 1700 rpm layered over 30% Percoll to isolate epithelial cells which were then used for FACS analysis.
In vivo BrdU labeling and annexin V staining: Day 7 or Day 10 after HCT, T cell proliferation was measured with a single intraperitoneal (i.p) injection of BrdU (2.5 mg/mouse, 100 mg/g) 3 hours before tissue harvesting. Day 21 after HCT, T cell proliferation was measured by three i.p. injections every 24 hours with BrdU (2.5 mg/mouse, 100 mg/g) beginning 72 hours before tissue harvesting. Analysis of donor CD8+ T cells for BrdU incorporation was performed according to the manufacturer's instructions (BD Pharmingen). For Annexin V staining, the percentage of Annexin V+ cells among donor CD8+ T cells was assessed by flow cytometry according to the manufacturer's instructions (eBioscience, San Diego, Calif.).
Real-time RT-PCR: Real-time RT-PCR analysis of mRNA for CCL25, CXCL9, CXCL10, CXCL11 was performed as described in the previous publication (1, 6). Primers used are as follows:
Relative expression levels of genes were normalized within each sample to the house keeping gene GAPDH.
Measurement of cytokines and liver function in serum: Cytokines in serum were measured by enzyme-linked immune sorbent assay (ELISA). The ELISA kits for IFN-γ, TNF-α and IL-2 were purchased from R&D Systems (Minneapolis, Minn.). ELISA kit for mouse IL-27 was purchased from Biolegend (San Diego, Calif.). Measurements of liver function (AST, ALT and ALB) were performed by the Charles River Clinical Pathology Laboratory (Wilmington, Mass.). Serum AST levels during GVL experiments was measured with Aspartate Aminotransferase activity assay kit purchased from abcam (Cambridge, Mass.).
Histopathology: Tissue specimens were fixed in formalin before embedding in paraffin blocks, sectioned and stained with H&E. Slides were examined at 200× or 400× magnification and visualized with an Olympus and a Pixera (600 CL) cooled charge-coupled device camera (Pixera, Los Gatos, Calif.). Tissue damage was blindly assessed on a scoring system, as described previously (1, 2). In brief, skin GVHD was scored by tissue damage in the epidermis and dermis and by loss of subcutaneous fat; the maximum score is 9. Salivary GVHD was scored by mononuclear cell infiltration and follicular destruction; the maximum score is 8. Liver GVHD was scored by the severity of lymphocytic infiltrate, number of involved tracts and severity of liver cell necrosis; the maximum score is 9. Lung GVHD was scored by periluminal infiltrates, pneumonitis, and the severity of lung tissues damage; the maximum score is 9. Gut GVHD was scored by mononuclear cell infiltration and morphological aberrations (e.g. hyperplasia and crypt loss), with a maximum score of 8.
Histoimmunofluorescent staining of intestinal Paneth cells and epithelial cells as well as thymic epithelial cells: Small intestine and colon tissues were harvested, formalin-fixed and paraffin embedded. Small intestines were stained with rat-anti-mouse IL-22Rα antibody (R&D Systems) and polyclonal rabbit anti-human lysozyme (Dakocytomation). Colon tissues were stained with anti-cytokeratin-Pan (Sigma). Frozen thymic tissues were put in PFA over night at 4° C., then transferred to sucrose over night at 4° C. Forty-eight hours later, the samples were embedded in OCT gel, frozen on dry ice and stored at −80° C. Thymus were stained with anti-UEA-1 (Vector lab) for medulla epithelial cells and anti-Cytokeratin 8 (DSHB) for cortical epithelial cells.
TUNEL assay of hepatocyte apoptosis: Paraffin sections were stained with DAPI and TUNEL according to the manufacturer's instructions (Roche, Indianapolis, Ind.) and imaged with the use of an Olympus IX81 Automated Inverted Microscope. Images were taken with a 400× objective and analyzed using Image-Pro Premier.
Bioluminescent imaging: Mice were injected with luciferase+ BCL1 cells (BCL1/Luc+) i.p. and monitored for expansion of those cells using bioluminescent imaging. In vivo imaging of tumor growth has been previously described (7). Briefly, mice were injected with 200 μl firefly luciferin i.p. (Caliper Life Sciences, Hopkinton, Mass.), anesthetized, and imaged by using an IVIS100 (Xenogen) and AmiX (Spectral) imaging system. Data were analyzed using Igor Pro 4.09A software purchased from WaveMetrics (Lake Oswego, Oreg.) and Amiview software purchased from Spectral Instruments Imaging (New York, N.Y.).
Production of mouse B7H1-Fc: B7H1-Fc-expressing plasmid was a kind gift from Dr. Lieping Chen (Yale University School of Medicine). The DNA plasmid contained the coding sequence for the murine B7H1 extracellular domain that was fused with the CH2-CH3 region of human IgG1 heavy chain. B7H1-Fc fusion protein was expressed transiently in Chinese Hamster Ovary Suspension (CHO-S) cell line using Thermo Fisher Freestyle CHO expression system as manufacture protocol. The supernatant of the transiently transfected CHO-S was collected after 7 days and passed through the protein G agarose beads (GenScript) packed column that had been equilibrated in 1× PBS pH.7.4. B7H1-Fc bound protein was washed with 1×PBS pH7.4, eluted with 0.1 M Glycine pH2.5, dialyzed in 1×PBS pH 7.4 and concentrated into 1.0 mg/ml aliquots before freezing in −80° C. until further use.
This example demonstrates that temporary depletion of donor CD430 T cells immediately after HCT preserves strong GVL effects, while effectively preventing both acute and chronic GVHD in multiple models.
A previous study showed that sorted CD8+ T cells from C57BL/6 donors did not induce acute GVHD but they induced chronic GVHD in lethally irradiated BALB/c recipients, as indicated by histopathology in salivary glands, a prototypic target organ of chronic GVHD. Depletion of CD430 T cells by treatment with anti-CD4 mAb on days 15 and 30 prevented the development of chronic GVHD, as indicated by prevention of tissue damage in all GVHD target tissues, especially in the salivary gland (11). It is disclosed herein that 1) in vivo administration of anti-CD4 mAb on the day of HCT was more effective in depleting donor CD430 T cells as compared to ex vivo depletion of CD430 T cells, as judged by percentage and yield of donor CD430 T cells in the spleen of recipients at 7 days after HCT (
The impact of temporary in vivo CD4+ T cell depletion on GVL effects against BCL1 tumor cells was evaluated (35, 36). TBI-conditioned BALB/c recipients were injected with luciferase-transfected BCL1 cells (BCL1/Luc, 5×106/mouse) together with TCD-BM (2.5×106) alone or TCD-BM+spleen cells (5×106). Recipients given spleen cells were treated with anti-CD4 mAb or control rat IgG on days 0, 14, and 28 days after HCT. BCL1/Luc tumor cell-bearing recipients transplanted with TCD-BM alone all died with progressive tumor growth by 20 days after HCT (
Depletion of donor CD8+ T cells by anti-CD8 (
Leukemia and lymphoma cells also infiltrates liver tissues. In vivo BLI indicated that tumor load started to decrease by day 7 and disappeared by day 12 after HCT (
The GVL capacity of this regimen was tested by using “GVL-resistant” blast crisis-chronic myeloid leukemia (BC-CML) in C57BL/6 background. Murine BC-CML cells obtained from W. Shlomchik were generated by retroviral transfer of bcr-abl and NUP98/HOXA9 fusion cDNAs. Like human BC-CML, murine BC-CML was relatively GVL resistant. At certain cell doses, allogeneic CD8+ T cells were not able to rescue recipients inoculated with BC-CML cells, although identical numbers of CD8+ T cells rescued almost all recipients inoculated with same number of chronic-phase chronic myelogenous leukemia (CP-CML) cells (37).
A/J BM (10×106) and spleen cells (10×106) were transplanted into lethally irradiated (1100 cGy) C57BL/6 recipients (38). The recipients were challenged with an intravenous injection of BC-CML (20×103 cells/mouse) at the time of HCT (37). The tumor cells killed all ( 12/12) GVHD-free recipients given TCD-BM alone within 30 days, and moribund mice had high percentages of BC-CML cells in the spleen, liver and bone marrow (
In further experiments, donor spleen cells were increased to 20 and 40×106 and the anti-CD4 treatment was extended to day 60 after HCT. 37.5% ( 6/16) recipients given 20 ×106 donor spleen cells died with progressive tumor growth, 62.5% ( 10/16) survived for more than 100 days without detectable tumor cells (
Whether administration of depleting anti-human CD4 mAb could prevent GVHD and preserve GVL effects in a xenogeneic model of GVHD was tested (39). NSG mice without or with human B cell lymphoma Raji cells (1×106/mouse) were used for GVHD or GVL experiments. Healthy human PBMC (20×106) were injected i.p. into mice that were then treated with depleting anti-human CD4 (clone IT1208, 200 μg/mouse) or control IgG twice weekly for 4 weeks (40). 4 human PBMC donors were tested. For each donor, 16 NSG mice were used, 8 for GVHD experiments and 8 for GVL experiments. Within each experiment, 4 recipients were treated with control IgG and 4 were treated with anti-CD4.
Anti-CD4 treatment effectively prevented xenogeneic GVHD in experiments with 3 of the 4 donors, and the 12 GVHD-free anti-CD4-treated NSG recipients survived for more than 100 days after PBMC injection (
In GVL experiments, control recipients given Raji cells alone all died with progressive tumor growth by 35 days. NSG mice given Raji cells and human PBMC were treated with IgG or anti-CD4 mAb. All 12 GVHD-free anti-CD4-treated mice survived for more than 100 days after PBMC injection (P<0.01,
This example demonstrates that depletion of donor CD430 T cells immediately after HCT increased serum IFN-γ but decreased serum IL-2 concentrations.
In experiments with C57BL/6 donors and BALB/c recipients, how in vivo depletion of donor CD4+ T cells immediately after HCT prevented acute GVHD while preserving GVL effects was explored. High serum levels of IFN- and TNF-α have been associated with acute GVHD (41). Contrary to expectation, depletion of donor CD4+ T cells increased serum IFN-
concentrations approximately 3-fold at 7 days after HCT (p<0.001). Serum IL-2 concentrations decreased by ˜50% (p<0.05), and serum TNF-α concentrations showed no significant differences from baseline (
is attributable to expansion of donor CD8+ T cells in lymphoid tissues, because the number of IFN-
+CD8+ T cells in the spleen of anti-CD4-treated recipients was ˜3 fold higher than in rat IgG-treated recipients (p<0.001), although the percentage of IFN-
+ cells among CD8+ T cells was similar in the two groups (
-producing CD8+ T cells in lymphoid tissues.
This example demonstrates that depletion of donor CD430 T cells immediately after HCT increased the numbers of donor CD8+ T cells in lymphoid tissues.
Next, the effects of in vivo CD4+ T cell depletion on donor CD8+ T cell expansion and tissue distribution were kinetically evaluated. At 5 days after HCT, the numbers of H-2Kb+ donor-type CD8+ T cells in the spleen and MLN were lower in anti-CD4-treated recipients than in rat IgG-treated recipients (P<0.01,
This example demonstrates that depletion of donor CD430 T cells immediately after HCT decreased the numbers of donor CD8+ T cells in the intestine and lung but not in the liver.
At 5 days after HCT, only a few donor CD8+ T cells infiltrated the colon, lung and liver, with no difference between recipients treated with IgG or anti-CD4. From day 7 to day 28 after HCT, the numbers of donor CD8+ T cells in the colon were markedly lower in anti-CD4-treated recipients than in IgG-treated recipients (p<0.01,
It was previously reported that donor T cell infiltration of gut tissues is regulated by their expression of gut tissue-specific homing and chemokine receptors (α4β7, CCR9, CXCR3), and by tissue release of the corresponding chemokines (CCL25 and Cxcl9-11) (42-45). By 7 days after HCT, more than 92% of the donor-type CD8+ T cells expressed a CD44hiCD62lo effector phenotype in both rat-IgG-treated and anti-CD4-treated recipients, indicating that CD8+ T cells were fully activated in both groups. Although donor CD8+ T cell infiltration of intestinal tissues (i.e., colon) was markedly decreased in anti-CD4-treated recipients at 7 days after HCT (
This example demonstrates that depletion of donor CD430 T cells immediately after HCT augmented donor CD8+ T cell apoptosis in the intestine and anergy/exhaustion in the liver, but not in the spleen.
The mechanisms was explored, whereby anti-CD4-treated GVHD-free recipients had reduced numbers of donor CD8+ T cell numbers in the colon and similar or higher numbers in the liver, while having increased numbers of donor CD8+ T cells in the spleen, as shown in
Alloreactive T cell infiltration also plays a critical role in damage to the liver (3). Although the numbers of liver infiltrating CD8+ cells were markedly higher in anti-CD4-treated recipients than in control IgG-treated recipients on day 10 after HCT (
Therefore, the proliferation and apoptosis of donor CD8+ T cells in the spleen, liver and colon tissues at 7 and 10 days after HCT were compared. At day 7, in vivo BrdU labeling showed that donor CD8+ T cells had significantly faster proliferation in the spleen, liver, and intestine tissues in anti-CD4-treated recipients as compared to IgG-treated recipients (P<0.01,
To evaluate anergy and exhaustion of donor CD8+ T cells, the CD8+ T cell expression levels (mean fluorescent index, MFI) of the anergy/exhaustion-related markers including Grail, Tim-3 and IL-R7α were compared. As compared to IgG-treated recipients, the CD8+ T cells from the spleen of anti-CD4-treated recipients did not have significant change in their expression of Grail, Tim-3 or IL-7Rα on day 7 (
Eomes regulates CD8+ T differentiation (48). Eomes+T-bet+ CD8+ T cells are effector cells with strong cytolytic function, while Eomes+PD-1+ CD8+ T cells are terminally differentiated exhausted cells (49, 50). Therefore, the impact of depletion of CD4+ T cells on CD8+ T expression of Eomes, T-bet, and PD-1 in the spleen and liver at 7 and 10 days after HCT were evaluated. CD8+ T cells from the spleen and liver of anti-CD4-treated recipients had significant increase in percentages of Eomes+T-bet+ and Eomes+PD-1+ cells, as compared to control IgG-treated recipients at days 7 and 10 after HCT (P<0.01,
This example demonstrates that depletion of donor CD430 T cells immediately after HCT allowed host-tissue expression of PD-L1 to tolerize infiltrating donor CD8+ T cells in GVHD target tissues but not in lymphoid tissues.
PD-L1/PD-1 interaction leads to T cell anergy and exhaustion (24), and simultaneous PD-L1/PD-1 and PD-L1/CD80 interactions augment apoptosis of activated alloreactive CD4+ T cells immediately after HCT (31). Depletion of donor CD4+ T cells increased serum levels of IFN- (
induces tissue expression of PD-L1 in GVHD target tissues (27, 29). Although IL-27 upregulates PD-L1 expression (51), no difference in serum IL-27 concentrations in recipients with or without anti-CD4-treatment was observed (
production and tissue expression of PD-L1 contributed to prevention of GVHD in anti-CD4-treated recipients was tested first. Spleen cells (5×106) from IFN-
−/− and wild-type C57BL/6 donors were transplanted into lethally irradiated BALB/c recipients, as described above. Indeed, anti-CD4 treatment did not prevent acute GVHD mediated by transplants from IFN-
−/− donors. All of recipients showed severe diarrhea and weight-loss, and ˜80% ( 8/10) of the recipients died by 30 days after HCT (
−/− transplants all had significantly down-regulated expression of PD-L1 immediately after HCT (
production and tissue PD-L1 expression contribute to GVHD prevention by administration of anti-CD4 immediately after HCT.
Furthermore, it was found that elevation of IFN- in anti-CD4-treated BALB/c recipients given wild-type C57BL/6 transplants was associated with upregulation of host intestinal epithelial cell expression of PD-L1 (
In addition, the role of host-tissue PD-L1 on acute GVHD severity induced by sorted CD4+ or CD8+ T cells was directly tested. While 2.5 or 5×106 sorted CD8+ T cells induced very little evidence of acute GVHD, the same numbers of donor CD4+ T cells induced severe lethal acute GVHD, and all the recipients died within 10 days (P<0.01,
In further experiments, the effect of host tissue expression of PD-L1 on the proliferation, apoptosis and anergy/exhaustion of CD8+ T cells in the spleen, liver and colon tissues of anti-CD4-treated recipients on day 7 after HCT was evaluated. As compared to anti-CD4-treated WT recipients, anti-CD4-treated PD-L1−/− recipients had no changes in proliferation or apoptosis of donor CD8+ in the spleen (
The impact of host-tissue expression of PD-L1 on donor CD8+ T expansion in the spleen on day 7 after HCT was evaluated. The numbers of splenic mononuclear cells (MNC), T cells, and CD8+ T cells were higher in anti-CD4-treated WT recipients than in rat-IgG-treated WT recipients (p<0.05-0.001,
CD80 and PD-1 expression by CD8+ T cells in the spleen was higher in anti-CD4-treated WT recipients than in rat IgG-treated recipients (p<0.05-0.001,
On day 7 after anti-CD4 treatment, expression of PD-1 and IL7Rα by CD8+ T cells in the spleen was higher in PD-L1−/− recipients than in WT recipients (p<0.001,
These results indicate that host-tissue expression of PD-L1 augments the apoptosis of infiltrating CD8+ T cells in the liver and intestine but not in the spleen of anti-CD4-treated recipients.
Expression of CD80 and PD-1 by infiltrating CD8+ T cells was higher in anti-CD4-treated WT and PD-L1−/− recipients than in rat IgG-treated WT recipients (p<0.001,
These results demonstrate that recipient tissue expression of PD-L1 contributed to increase of apoptosis of colon infiltrating CD8+ T cells and prevention of intestinal GVHD after CD4+ T cell depletion.
The impact of host-tissue expression of PD-L1 on anergy and exhaustion of CD8+ T cells infiltrating the liver at day 7 after HCT was evaluated. Anergic CD8+ T cells upregulate expression of GRAIL and down-regulate expression of IL-7Rα without significant changes in TIM3 expression, while exhausted CD8+ T cells express high levels of both PD-1 and TIM3 (72-75). Anergic and exhausted T cells gradually lose proliferative capacity and effector function (e.g., production of IFN-γ) (72, 73). As compared to rat-IgG-treated recipients, liver infiltrating CD8+ T cells of anti-CD4-treated recipients expressed higher levels of CD80, PD-1, and GRAIL (p<0.01), lower levels of IL-7Rα (p<0.01), and similar levels of TIM3, (
These results suggest that the infiltrating CD8+ T cells in the liver of anti-CD4-treated recipients immediately after HCT were becoming anergic.
In the absence of recipient PD-L1, expression of GRAIL by CD8+ T cells was not significantly upregulated, and expression of IL-7Rα was not down-regulated after anti-CD4 treatment (
At 21 days after HCT, CD8+ T cells infiltrating the liver were exhausted in anti-CD4-treated recipients but not in rat-IgG-treated recipients, as judged by their up-regulation of PD-1 and TIM-3 (p<0.01,
Taken together, these results show that CD8+ T cells infiltrating the liver in anti-CD4-treated recipients immediately after HCT became progressively anergic and exhausted through mechanisms dependent on expression of PD-L1 in the recipient.
The expression levels (MFI) of Grail, Tim-3, IL-7Rα and percentage of Eomes+T-bet+CD8+ and Eomes+PD-1+CD8+ T cells in the spleen and liver of PD-L1−/− recipients and controls at 7 days after HCT were compared. The absence of host-tissue expression of PD-L1 did not significantly change donor CD8+ T expression of Grail or Tim-3 in the spleen, although expression of IL-7Rα was higher in PD-L1−/− recipients than in WT recipients (
Furthermore, it was found that human T cells could interact with mouse PD-L1 (
This example demonstrates that depletion of donor CD430 T cells immediately after HCT led to donor CD8+ T cell upregulated expression of PD-L1 and CD80 in lymphoid tissues, which preserved GVL effects.
Since donor T cell expression of PD-L1 augments acute GVHD lethality in recipients transplanted with both CD4+ and CD8+ T cells (30), the effect of donor CD8+ T expression of PD-L1 in the expansion and GVL activity of CD8+ T cells in GVHD-free anti-CD4-treated recipients was evaluated. Anti-CD4-treatment significantly upregulated CD8+ T cell expression of PD-L1 in the spleen and liver but not in the colon (
Transplantation of sorted Thy1.2+ T cells from PD-L1−/− C57BL/6 donors and TCD-BM cells from wild-type C57BL/6 donors led to marked reduction of donor CD8+ T expansion in the spleen of anti-CD4-treated recipients immediately after HCT, as compared to Thy1.2+ T cells from WT donors (P<0.001,
To further evaluate the role of PD-L1/CD80 interaction on CD8+ T cell survival and expansion, an anti-PD-L1 mAb (43H12) that specifically blocks PD-L1/CD80 interaction without interfering with PD-L1/PD-1 interaction was used (26). The 43H12 mAb was injected i.p. into anti-CD4-treated WT recipients on days 0 and 2 after HCT. As compared to control IgG treatment, blockade of PD-L1/CD80 interaction also markedly decreased donor CD8+ T cell expansion in the spleen. This finding was associated with augmented apoptosis, reduced expression of BCL-XL, and increased percentage of Eomes+PD-1+ cells (
Finally, the impact of PD-L1/CD80 interaction on GVL activity in anti-CD4-treated recipients was evaluated. Since BCL1/luc+ tumor cells in anti-CD4-treated recipients were eliminated within 12 days after HCT without relapse by 100 days after HCT (
This example demonstrates that depletion of donor CD430 T cells immediately after HCT augmented thymic infiltrating CD8+ T cell anergy.
The impact of host-tissue expression of PD-L1 on donor CD8+ T cell expansion in the thymus was evaluated. On day 7 after HCT, the number of thymic mononuclear cells was higher in anti-CD4 treated WT recipients than in rat IgG-treated WT recipients (p<0.01,
These results indicate that, in the absence of donor CD430 T cells, thymic infiltrating CD8+ T cell interaction with host tissue PD-L1 via CD80 and PD-1 lead to donor CD8+ T cell proliferation and development of anergy, such that the accumulation of infiltrating CD8+ T cells did not cause thymic tissue damage.
This example demonstrates that injection of an anti-IL-2 mAb after HCT prevented acute GVHD in BALB/c recipients with C57BL/6 transplants.
As shown in
As stated above, the foregoing are merely intended to illustrate the various embodiments of the present invention. As such, the specific modifications discussed above are not to be construed as limitations on the scope of the invention. It will be apparent to one skilled in the art that various equivalents, changes, and modifications may be made without departing from the scope of the invention, and it is understood that such equivalent embodiments are to be included herein. All references cited herein are incorporated by reference as if fully set forth herein.
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The present application is a continuation of International Application No. PCT/US2018/019524, filed Feb. 23, 2018 which claims the benefit of U.S. Provisional Application No. 62/462,853, filed Feb. 23, 2017, both of which are incorporated herein by reference in their entireties.
The present invention was made with government support under Grant No. R01 AI066008, 2R56AI66008-11, RO1 AI095239, and P30CA033572, awarded by the National Institutes of Health (NIH). The Government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 62462853 | Feb 2017 | US |
| Number | Date | Country | |
|---|---|---|---|
| Parent | PCT/US2018/019524 | Feb 2018 | US |
| Child | 16543472 | US |
