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IMMUNOBIOLOGY
From the Departments of Haematological Medicine and
Histopathology, Guy's, King's and Thomas' School of Medicine,
Denmark Hill, London, England.
CAMPATH antibodies recognize CD52, a phosphatidylinositol-linked
membrane protein expressed by mature lymphocytes and monocytes. Since
some antigen-presenting dendritic cells (DCs) differentiate from a
monocytic progenitor, we investigated the expression of CD52 on
dendritic cell subsets. Four-color staining for lineage markers (CD3,
14, 16, 19, 20, 34, and 56), HLA-DR, CD52, and CD123 or CD11c
demonstrated that myeloid peripheral blood (PB) DCs, defined as
lineage The CAMPATH series of monoclonal antibodies
recognize CD52, a phosphatidylinositol-linked antigen that is expressed
by most mature lymphocytes and monocytes.1 CAMPATH-1M and
1G are rat IgM and IgG antibodies that have been used in vitro and in
previous phase 1/2 clinical trials. A humanized form of the antibody,
CAMPATH-1H (approved name Alemtuzumab), has recently been approved for
refractory B-cell chronic lymphocytic leukemia. Several properties of
the CD52 antigen make it a particularly good therapeutic target. These include the high antigen density, lack of internalization, and the
proximity of the antibody binding site to the cell surface, which may
contribute to the efficiency of complement-mediated cytotoxicity that
characterizes CAMPATH antibodies.2 Administration of
CAMPATH antibodies results in the rapid clearance of normal and
malignant T and B lymphocytes from the circulation, a finding that has
led to its use as an immune suppressant3 and in the treatment of mature T and B lymphoproliferative
disorders.4,5
CAMPATH antibodies have also proved to be a convenient and efficient
means of achieving T-cell depletion in patients undergoing allogeneic
hematopoietic stem cell transplantation (allo-HSCT).6,7 Conditioning regimens that incorporate CAMPATH antibodies have been
associated with a low incidence of graft-versus-host disease (GvHD) and
rejection.8-11 Although these effects can be explained by
depletion of donor and recipient T cells, we investigated the effects
of CAMPATH antibodies on antigen-presenting dendritic cells (DCs),
which have recently been shown to have a role in the initiation of
GvHD12 and, by virtue of their monocytic
lineage,13 might also express CD52.
CD52 expression by blood and tissue DCs
Phenotypic studies.
Measurements of peripheral blood (PB) DC numbers were performed on
EDTA-anticoagulated whole blood from healthy volunteers and
patients receiving alemtuzumab (Millenium-Ilex, Guildford, United
Kingdom) as therapy for lymphoproliferative disorders or as
conditioning treatment for reduced-intensity allo-HSCT. Patients with
lymphoproliferative disorders received 30 mg of intravenous alemtuzumab on alternate days and allo-HSCT patients received 20 mg
daily for 5 consecutive days. The studies were approved by the local
research and ethics committee and patients gave informed consent. No
concurrent cytotoxic drugs were administered to the patients under
study. Steady-state numbers of PBDCs were measured in 12 healthy volunteers and in 5 patients prior to and on day 5 of
treatment with alemtuzumab.
Functional studies.
Cell purification. For the functional assays of antigen
presentation, peripheral blood from healthy volunteers was
taken into 10 U/mL preservative-free heparin and mononuclear
cells (PBMCs), prepared by Ficol Hypaque centrifugation at
400g for 20 minutes at room temperature, followed by 2 washes with phosphate-buffered saline. Further cell purification was
performed with immunomagnetic beads used according to the
manufacturer's instructions. Purification of responder T cells for the
allogeneic mixed lymphocyte reactions (allo-MLRs) was performed by
negative depletion of CD14, HLA-DR/DP, and CD16 and CD56+
cells from PBMCs (Dynal, Oslo, Norway). For the autologous presentation assays, CD4+ T cells were isolated with the above cocktail
with the addition of anti-CD8. CD52 depletion was performed with
CAMPATH-1G (Therapeutic Antibody Centre) and sheep antirat conjugated
beads (Dynal). Cell purity was confirmed by flow cytometry and ranged
from 85% to 97%. In some experiments, further depletion of
CD123+ cells from the CD52 Allogeneic mixed lymphocyte reaction. Purified T lymphocytes were cocultured for 5 days at 105 per well in RPMI/10% fetal calf serum (FCS) at 37°C and 5% CO2, in triplicate, with irradiated (35 Gy) allogeneic stimulator cells comprising either unfractionated PBMCs or the same number of antibody-depleted PBMCs at the stated concentrations. In some experiments, purified monocytes, T cells, and B cells were added back to CD52-depleted cells in the same proportion in which they had been present prior to depletion. This was achieved by selecting from the same volume of PBMCs used to arrive at the stated number of CD52-depleted cells. On day 5, 3H-thymidine (0.0185 Bq [0.5 µCi] per well; Amersham International, Little Chalfont, United Kingdom) was added for the final 20 hours. The cells were then harvested and 3H-thymidine incorporation was measured by liquid scintillation. The significance of differences in response to PBMCs and CD52-depleted cells was assessed by Student t test. Primary antigen-specific responses. To assess presentation of primary antigen by CD52-expressing cells, we used irradiated normal PBMCs or the same number of antibody-depleted cells as stimulators and purified autologous CD4 cells as responders in RPMI 1640 medium supplemented with 10% heat-inactivated autologous serum. Keyhole limpet hemocyanin (KLH; Sigma, Poole, United Kingdom) was added where indicated at a concentration of 25 µg/mL. Add-back of CD3-, CD14-, and CD19-expressing cells was achieved as outlined above. 3H-thymidine was added on day 7 for the final 20 hours and the cells were harvested and analyzed as previously described. CD52 expression by monocyte-derived DCs Monocytes were enriched from PBMCs by plastic adherence in RPMI + 1% FCS for 2 hours at 37°C. After removal of nonadherent cells, the adherent cells were washed and then incubated in RPMI + 10% FCS with 100 ng/mL GM-CSF (Novartis, Camberley, United Kingdom) and 500 U/mL interleukin-4 (IL-4) (R&D Systems, Abingdon, United Kingdom) for 6 days. Maturation was achieved by the addition of lipopolysaccharide (LPS; Sigma), 100 ng/mL, to an aliquot of the cells on day 4. Differentiation to DCs was assessed by staining with CD1a, CD14, CD80, and CD83 (BD Biosciences) and CD52 (Serotec, Oxford, United Kingdom) and analysis by flow cytometry. The significance of changes in the percentage of cells expressing CD52 was assessed by paired t test.
Peripheral blood dendritic cells express CD52 Cells of the monocyte-macrophage lineage share a common progenitor with some types of DC and are known to express CD52. We therefore investigated whether peripheral blood DCs, defined by the lack of lineage markers and strong expression of HLA-DR, also express CD52. Analysis by 4-color flow cytometry revealed that CD11c+ "myeloid" DCs uniformly express CD52 (Figure 1A), while CD52+ and CD52 populations of CD123+ lymphoid DCs were
evident (Figure 1B). A population of
linHLA-DR+CD52+ cells negative
for both CD11c and CD123 was also suggested by these experiments, and
this was confirmed by the addition of CD123 and CD11c to the lineage
cocktail and staining for CD52 and HLA-DR (data not shown).
To determine whether the CD52+ DCs identified by phenotypic
criteria have the appropriate functional properties, we assessed the
effect of removing CD52+ cells from the stimulatory arm of
the allo-MLR. CD52
To confirm the antigen-presenting capabilities of CD52-expressing
cells, we studied the effect of CD52 depletion on the presentation of
the potent soluble neoantigen KLH to purified autologous CD4 T
lymphocytes. In each of 3 experiments with different donors, CD52
depletion significantly inhibited the primary immune response to KLH
(Figure 3A). Add-back of purified
CD3+, CD14+, and CD19+ cells
failed to reconstitute antigen-presenting ability, and lineage-depleted PBMCs showed enhanced activity compared with CD52-depleted cells, suggesting that CD52-expressing DCs and not cells
of another type were responsible for the presentation of KLH (Figure
3B). Depletion of the CD52
Peripheral blood dendritic cells are depleted during treatment with alemtuzumab We next studied the effect of treatment with alemtuzumab on the number of circulating linHLA-DR+ DCs in a
group of 5 patients being treated with alemtuzumab and no other agents.
Previous studies had shown the normal range for PBDCs to be 21± 2.6 (SEM)/µL (n = 12) . DC numbers in the patients studied
were within this range prior to therapy. The administration of
alemtuzumab resulted in a significant (P = .043)
reduction in circulating DCs in all patients (Figure
4). As expected, a fall in the
circulating number of lymphoid cells, but not neutrophils, was also
noted. This is in keeping with the tissue distribution of CD52, which
is expressed by mature T and B lymphocytes but not neutrophils.
Expression of CD52 by monocyte-derived DCs and by tissue DCs in vivo Since the effect of alemtuzumab therapy on immune responses in vivo will depend on whether CD52 is expressed by immature DCs in the tissues and mature antigen-primed DCs, we studied CD52 expression by monocyte-derived DCs (MoDCs) in vitro and by DCs in the normal epidermis and intestinal mucosa in vivo. DCs derived by differentiation of monocytes in GM-CSF and IL-4 showed the expected gain in CD1a and loss of CD14 expression. The addition of LPS resulted in down-regulation of CD1a and up-regulation of CD83, compatible with activation-induced maturation (data not shown). During the culture period there was a progressive fall in the proportion of cells expressing CD52 when gating was performed on the DC population. A similar loss of CD52 expression was also observed in the lymphocytic fraction, suggesting that it may be a phenomenon associated with the duration of time in tissue culture. The addition of LPS resulted in significant further loss of CD52 by MoDCs (P = .013, n = 4) but had no effect on CD52 expression by lymphocytes (Figure 5).
Staining of adjacent tissue sections with S100 and CD52 showed that
epidermal Langerhans cells and small-intestine DCs do not express CD52
(Figure 6). The presence of melanocytes
at the epidermal/dermal junction and neuronal fibrils in the submucosa of the intestine served as positive internal controls for the S100
antibody. CD52 staining was validated by the presence of lymphocytes in
both tissues.
Normal peripheral blood contains at least 2 DC subsets that together make up between 0.01% and 0.1% of circulating white blood cells.15,16 These include myeloid DCs defined by the absence of lineage markers and strong expression of HLA-DR, CD11c, and CD33, and a lymphoid subset that expresses the IL-3 receptor CD123 instead of CD11c or CD33. Both are strong stimulators of the allo-MLR, but only the myeloid subset, thought to be the precursor of the cutaneous Langerhans cell,17 can process soluble antigens.16 The results presented here indicate that most CD11c+ myeloid PBDCs and some CD123+ lymphoid DCs express CD52, the antigen recognized by CAMPATH antibodies. This was confirmed by functional experiments showing that the removal of CD52-expressing cells reduced both the allostimulatory activity of PBMCs and the ability to present the potent soluble neoantigen KLH to autologous T cells. That DCs, rather than another CD52-expressing cell, were responsible
for the activity in these assays was confirmed by lineage-depletion studies and by add-back of purified monocytes and lymphocytes. Lineage
depletion, a maneuver known to enrich DCs, augmented both the allo-MLR
and the KLH-presenting activity, while add-back of monocytes and
lymphocytes variably reconstituted the allo-MLR but had no effect on
the primary antigen presentation assay. Further depletion of
CD123+ cells, which would additionally remove
CD52 As anticipated from the observation that most PBDCs express CD52, the
administration of intravenous alemtuzumab had marked effects on the
numbers of peripheral blood lineage Our data also suggest that the expression of CD52 is a feature of an immature stage of DC development, since in vitro differentiation from monocytes in GM-CSF and IL-4 followed by activation-induced maturation in LPS was associated with progressive loss of the antigen. In addition, as recently noted by others,18 DCs in the epidermis (Langerhans cells) do not express CD52, and we now show that this is also true of DCs in the small-intestine mucosa. These findings raise questions about the role of circulating DCs, which have previously been proposed as the precursors of tissue DCs such as Langerhans cells.17 Either migration from blood to tissues is associated with loss of CD52, or DCs in these tissues must arise from another cell type. If the former is true, then, depending on the rate of turnover of DCs in that particular site, therapy with alemtuzumab should eventually reduce tissue DCs by depleting their circulating precursors. In the latter case, therapy with alemtuzumab would not affect DC numbers in tissues such as the skin and gut. Sequential studies of blood and tissue DC numbers in patients receiving long-term alemtuzumab therapy will be required to clarify this point. These findings have important implications for the clinical use of alemtuzumab. Since their introduction during the 1980s, CAMPATH antibodies have been used as an immunosuppressive agent in patients with autoimmune disorders such as rheumatoid arthritis3 and multiple sclerosis19 and in solid organ20 as well as bone marrow transplantation.8-11 It is particularly useful in the treatment of T- and B-cell malignancies,5,21 for which a product license was recently granted. In the context of malignant disease, in which there is frequently pre-existing immunosuppression due to disease and other therapy, a relatively high incidence of opportunistic infection has been noted. While this is undoubtedly due in large measure to the depletion of T and B lymphocytes, any effect on antigen-presenting function could significantly compound this effect. Since CD52 is expressed by blood but not tissue DCs and is lost following activation-induced maturation, it seems likely that at least in the short term, antigen-presenting function will remain intact during alemtuzumab therapy. In particular, antigen-primed DCs that have matured and migrated to lymphoid tissue should be spared. The effect of depletion of PBDCs by alemtuzumab is less certain, since their role in immune function has not been defined. If this population of DCs represents the precursors of tissue DCs, then as noted above, depending on the turnover rate, alemtuzumab therapy should eventually result in impaired antigen-presenting function. Tissue depletion might be accelerated by damaging treatment modalities such as high-dose chemotherapy or radiation, since cellular injury and inflammatory stimuli are known to cause migration of DCs from the periphery into lymphoid tissue. If, on the other hand, the function of PBDCs is to process and present circulating pathogens, then alemtuzumab therapy will have rapid deleterious effects on this aspect of immune function. CAMPATH antibodies have proved to be particularly effective when incorporated into conditioning regimens for allo-HSCT, and the findings reported here are of particular relevance in this context. Several strategies have been employed, including in vivo administration prior to and after infusion of the donor stem cells or in vitro as a means of reducing the infused T- and B-cell dose. Although never formally evaluated in randomized trials, regimens that incorporate CAMPATH antibodies have been notable for their low incidence of graft failure and GvHD.9 The risk of posttransplantation lymphoma has also been low, presumably because of depletion of donor B cells.22 Interestingly, the risk of GvHD remains low when alemtuzumab administration is confined to the pretransplant period.10,11,23 This finding has been ascribed to in vivo depletion of donor T cells by antibody carried over from the pretransplantation period. When used "in the bag" as an in vitro T-cell depletion strategy, alemtuzumab should also deplete myeloid DCs, which may reduce the risk of rejection by removing the ability to present donor histocompatibility antigens to naive host T cells that have survived conditioning therapy. As discussed above, the effects of alemtuzumab therapy on DC populations and antigen presentation in vivo are unclear because of the lack of CD52 expression by tissue DCs and the uncertain relationship between these cells and DCs in the blood. Any effect of alemtuzumab on tissue DCs is, as noted earlier, likely to be influenced by other factors, such as tissue damage due to concurrent chemo- or radiotherapy or infection. In the context of allo-HSCT, it will be important to fully define the effects of alemtuzumab on DC function in vivo, as there is a persuasive theoretical and experimental basis for the role of these cells in the induction of GvHD. Although T lymphocytes can recognize major histocompatibility complex (MHC) disparity without the need for specialized antigen presentation,24 this does not apply to unprimed MHC identical sibling transplants, where GvHD and rejection presumably require the processing and presentation of minor histocompatibility antigens by professional antigen-presenting cells. This is borne out by murine allo-HSCT experiments in which the presence of functional host DCs was required for the induction of GvHD.12 It may be possible, therefore, to design improved conditioning regimens exploiting the effects of alemtuzumab on antigen presentation. In summary, we have shown that peripheral blood cells with the phenotype and function of myeloid dendritic cells express CD52 and are depleted by therapy with alemtuzumab. This is unlikely to have effects on antigen-presenting function in the short term, however, as tissue DCs and mature MoDCs do not express CD52 and are therefore unlikely to be depleted. Prolonged administration of alemtuzumab or concurrent tissue damage may reduce the number of DCs in the tissues by depleting their putative peripheral blood precursors; however, confirmation of this will require serial studies of tissue DC numbers.
We thank Professors Geoffrey Hale and Herman Waldmann of the University of Oxford for helpful advice and the supply of reagents for this study.
Submitted August 13, 2001; accepted April 4, 2002.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Stephen Devereux, Department of Haematological Medicine, Kings College Hospital, London SE5 9RS, England; e-mail: stephen.devereux{at}kcl.ac.uk.
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© 2002 by The American Society of Hematology.
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Top
Abstract
Introduction
prophylactic use after kidney transplantation. A randomized controlled clinical trial.
Transplantation.
1989;48:248-253