A R T I C L E S

Historically, ABO compatibility between donor and recipient has beenrequired for successful organ transplantation. Transplantation of ABO-incompatible donor organs into recipients with preformed antibodies todonor A or B antigens usually results in ‘hyperacute’ or acute antibody-mediated rejection, initiated by antibody binding to graft antigensresulting in complement activation and thrombosis of graft vasculature.Transplantation of ABO-incompatible kidney and liver grafts has beenaccomplished with some success, but aggressive immunosuppressivestrategies are required in mature individuals, often including splenec-tomy1–6. ABO-incompatible heart transplantation has been performedrarely, usually unintentionally, because of the high risk of lethal anti-body-mediated rejection5.

In 1945, Owen defined the inherent susceptibility to immune toler-ance induction during immaturity as a consequence of antigen exposureduring gestation7. He reported that dizygotic twin calves exposed toallogeneic blood from shared placental circulation did not reject skingrafts from each other. Burnet then linked immune tolerance to devel-opmental events8. Medawar and colleagues showed that tolerance couldbe induced intentionally (‘acquired’ tolerance), whereby introduction ofantigens to fetal or neonatal mice resulted in permanent and specificabrogation of the development of immune responsiveness to those anti-gens9. Evidence emerged showing several mechanisms of neonatallyinduced tolerance, including functional inactivation (anergy) or dele-tion of immature alloreactive clones, suppression of alloreactivity byimmunoregulatory cells and, in B cells, receptor editing10–16. Althoughstudied extensively at both T-cell and B-cell levels in mouse models,acquired neonatal tolerance has not been previously shown in humans.

Neonatal tolerance was not generally viewed as a clinically relevantphenomenon because human immune maturation at birth is consid-erably more advanced than in mice, thus presumably beyond suscepti-bility to tolerance induction17,18. Nonetheless, the human infantmanifests several aspects of immunologic immaturity, notably defi-ciency of humoral responsiveness to stimulation by carbohydrate (‘T-independent type 2’) antigens19–21. This includes development ofisohemagglutinins or ‘natural’ antibodies to nonself A and B bloodgroup antigens, which remain low during the first months of life22,23.Reasoning that humoral rejection of ABO-incompatible heart graftswould not occur in the absence of preformed donor-specific antibody,we undertook a trial of ABO-incompatible heart transplantation ininfants, using standard immunosuppression without splenectomy24.There were no cases of hyperacute or acute humoral rejection, norclinical problems attributable to blood group incompatibility. We havenow studied the B-cell immunobiology of ABO-incompatible infantheart transplant recipients and, for the first time, found evidence ofneonatally acquired donor-specific B-cell tolerance in humans.

RESULTSOntogeny of blood group–specific antibodiesWe analyzed sequential serum samples from 13 recipients of ABO-incompatible grafts and 3 recipients of ABO-compatible grafts todetect circulating A-specific and B-specific antibodies before trans-plantation.

Whereas serum A-specific antibodies developed normally ingroup O recipients of heart grafts from group B donors (B→O

1Infection, Immunity, Injury and Repair Program and 2Division of Cardiology, Department of Pediatrics, The Hospital for Sick Children, 555 University Avenue,Toronto, Ontario M5G1X8, Canada. 3Department of Pathology, University of Miami School of Medicine, Jackson Memorial Hospital, 1611 NW 12 Avenue, Miami,Florida 33136, USA. 4Department of Pediatric Laboratory Medicine, The Hospital for Sick Children, 555 University Avenue, Toronto, Ontario M5G1X8, Canada.5Department of Transplantation Biology, Mayo Clinic, Medical Sciences 2-66, 200 First Street SW, Rochester, Minnesota 55905, USA. Correspondence should beaddressed to L.J.W. (lori.west@sickkids.ca).

Published online 24 October 2004; doi:10.1038/nm1126

Donor-specific B-cell tolerance after ABO-incompatibleinfant heart transplantationXiaohu Fan1, Andrew Ang1, Stacey M Pollock-BarZiv2, Anne I Dipchand2, Phillip Ruiz3, Gregory Wilson4,Jeffrey L Platt5 & Lori J West1,2

Although over 50 years have passed since its first laboratory description, intentional induction of immune tolerance to foreignantigens has remained an elusive clinical goal. We previously reported that the requirement for ABO compatibility in hearttransplantation is not applicable to infants. Here, we show that ABO-incompatible heart transplantation during infancy results in development of B-cell tolerance to donor blood group A and B antigens. This mimics animal models of neonatal tolerance andindicates that the human infant is susceptible to intentional tolerance induction. Tolerance in this setting occurs by elimination ofdonor-reactive B lymphocytes and may be dependent upon persistence of some degree of antigen expression. These findingssuggest that intentional exposure to nonself A and B antigens may prolong the window of opportunity for ABO-incompatibletransplantation, and have profound implications for clinical research on tolerance induction to T-independent antigens relevant toxenotransplantation.

NATURE MEDICINE VOLUME 10 | NUMBER 11 | NOVEMBER 2004 1227

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

recipients; Table 1 and Fig. 1a) despite nonspecific immunosup-pressants, we noted a persistent deficiency in circulating B-specificantibodies (Fig. 1b). Similarly, we observed a deficiency in devel-opment of A-specific antibodies in A→O recipients (patients 5–9),whereas B-specific antibodies developed normally (Fig. 1c,d).Serum A-specific and B-specific antibodies remained negligible inB→A (patients 11 and 13) and A→B (patient 12) recipients (Fig. 1e,f), whereas O→O recipients (patients 14–16) developedhigh titers of both A-specific and B-specific antibodies (Fig. 1g,h),similar to nonimmunosuppressed individuals. This selective deficiency in isohemagglutinin ontogeny suggests that donor-specific B-cell tolerance has occurred after ABO-incompatibletransplantation, induced by exposure to donor blood group antigens during the ill-defined period of immunologic immaturityin the human infant.

Analysis of successful ABO-incompatible adult human kidneytransplants showed the phenomenon of graft accommodation,whereby grafts survived without damage despite intragraft deposi-tion of donor-specific antibodies and complement components suchas C4d3,4. Thus, low levels of circulating antibodies may reflect depo-sition within the graft rather than impaired production. To confirmthat deficient serum donor-specific antibodies in ABO-incompatiblegraft recipients was not a result of intragraft deposition, we assessedendomyocardial biopsies for immunoglobulin and complement deposition in addition to routine analysis for acute rejection. Insequential biopsies obtained during follow-up ranging from 1.5 to 5years after transplant, immunohistochemistry staining showed nodeposition of immunoglobulin (IgG or IgM) or complement (C3 orC4d, or both); (Table 1 and Fig. 2a,b; one representative example is shown of a negative biopsy; comparison is made to a case ofantibody-mediated rejection resulting from HLA sensitization). Wedid not observe any other histologic evidence of antibody-mediatedgraft damage attributable to ABO incompatibility. Thus, lack of

circulating and intragraft antibodies confirmed impairment ofantibody production against donor A and B antigens. (Note: As previously reported, the incidence of acute cellular rejection in the original cohort was slightly less in ABO-incompatible graft recipients than in ABO-compatible graft recipients24; chronic vascular rejection manifested as accelerated graft vasculopathydeveloped in one ABO-incompatible recipient (Table 1)).

Patients 4 and 10 (both AB→O) demonstrated divergent patterns of A-specific antibody development. Although bothpatients did not produce clinically significant levels of serum B-specific antibodies (Fig. 1b,f), only patient 10 remained deficientin A-specific antibody (Fig. 1a,e). Notably, patient 10 received agroup A1B graft, while patient 4’s donor was group A2B. Theuncommon A2 subtype is characterized by low-density cell-surfaceA antigen expression compared to the prevalent A1 subtype25, andgroup A2B cells have even fewer A antigen sites than A2 cells26. Inthis setting, exposure to A2B graft antigens would seem to inducetolerance only to the B antigens, whereas A1B antigen exposurewas tolerogenic for both A and B antigens.

Absence of agglutination-inhibiting factorsThe apparent deficiency of donor-specific isohemagglutinins insera from ABO-incompatible graft recipients (Fig. 1) could resultfrom interference with donor-specific antibodies by serum factors,such as idiotype-specific antibodies. To assess this possibility, wedevised a modified hemagglutination assay in which serum from agroup O volunteer was serially diluted using patients’ sera ratherthan saline before incubation with group A or B erythrocytes.Titers at which agglutination of group A and B erythrocytesoccurred by group O serum diluted with patients’ sera were equalto titers obtained with saline dilution of the same group O serum,that is, A-specific >1:256 and B-specific >1:256. Sera from all 13ABO-incompatible recipients showed no inhibition of erythrocyte

1228 VOLUME 10 | NUMBER 11 | NOVEMBER 2004 NATURE MEDICINE

a b c d

e f g h

Figure 1 Development of serum A-specific and B-specific antibodies in recipients of ABO-incompatible and ABO-compatible heart transplants showing a persistent and selective deficiency in donor-specific antibody. Isohemagglutinin titer values indicate the inverse of the highest dilution of serum giving a positive result. Transplantation was performed at (a,b) A-specific (a) and B-specific (b) antibody development in B→O and A2B→O recipients (n = 4).(c,d) A-specific (c) and B-specific (d) antibody development in A→O recipients (n = 5). (e,f) A-specific (e) and B-specific (f) antibody development inA→B, B→A and A1B→O recipients (n = 4). (g,h) A-specific (g) and B-specific (h) antibody development in O→O recipients (n = 3).

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

agglutination by serially diluted serum from a single group O volunteer. Thus, even at dilutions of 1:256 (group O serum:patientserum) patients’ sera were incapable of preventing erythrocyteagglutination by group O serum. This supports our conclusionthat donor-specific isohemagglutination deficiency in recipientsof ABO-incompatible transplants results from absence of donor-specific antibody.

Donor antigen persistenceIn animal models, persistence of donor antigen has been shown tobe important for induction of donor-specific tolerance27,28. Usingimmunoperoxidase staining, we assessed endomyocardial biopsyspecimens for A and B antigen expression. We analyzed at least twosequential biopsies from seven recipients of ABO-incompatiblegrafts and two recipients of ABO-compatible grafts; we showedexpression of A or B antigen, or both, of the heart donor in allcases that were not group O (Table 1 and Fig. 2). A antigen expres-sion is observed in a graft biopsy from a group A donor 2 yearsafter transplantation into a group O recipient (Fig. 2c,d); likewise,persistent B antigen expression is apparent in a biopsy from agroup B donor 4 years after transplantation into a group O recipient (Fig. 2e,f).

Patient 16, a group O individual, required urgent retransplantation(unrelated to ABO status) 3 d after primary ABO-incompatibletransplantation (group B donor), receiving an ABO-compatiblesecond graft (group O donor). This patient developed A-specificand B-specific antibodies in a normal pattern (Fig. 1g,h), suggest-ing that only transient exposure to donor antigens was insufficientto induce donor-specific antibody deficiency.

Assessment of B-cell functionTo ascertain cellular mechanisms of donor-specific antibody defi-ciency, we performed B-cell functional and phenotypic analyses

using whole blood samples from a subset of 12 ABO-incompatibleand 3 ABO-compatible graft recipients. Anergic lymphocytes havebeen defined as antigen-specific, nonresponsive cells that canrecover responsiveness after in vitro culture with appropriate stim-ulation29,30. The inability of mouse neonatal B lymphocytes torespond to antigens can be overcome with cytokine stimula-tion31,32, and antibody production by human neonatal lympho-cytes in response to CD3-specific stimulation can be elicited withinterleukin (IL)-2 and IL-4 (ref. 33).

To determine whether tolerance in infant recipients of ABO-incompatible grafts resulted from functionally inactive donor-spe-cific B cells, we cultured patient-derived peripheral bloodmononuclear cells (PBMC) in vitro with nonspecific cytokine orspecific donor antigen stimulation, and measured specific anti-body levels in culture supernatants by ELISA. Results were consis-tent amongst all patients tested (Table 1 and Fig. 3). A-specific IgMproduction by PBMC from two A→O recipients cultured withoutstimulation was low, comparable to PBMC from a group A con-trol, whereas cultured PBMC from an O→O recipient exhibitedmodest A-specific antibody production (Fig. 3a). When culturedwith IL-2 and IL-4, A-specific IgM production was markedlyincreased by PBMC from the O→O recipient, but remained mini-mal in cultures from the A→O recipients and the group A control(Fig. 3b). We obtained similar results when we cultured cells withgroup A erythrocytes (Fig. 3c). To evaluate the functional capacityof patients’ PBMC under in vitro culture conditions, we measuredantibody production to the ‘third-party’ carbohydrate epitopeGal-α1-3β1-4GlcNAc-R (α-gal), a major xenoantigen againstwhich humans produce ‘natural’ antibodies34. Production of α-gal-specific IgM was not substantially different amongst cytokine-stimulated culture groups (Fig. 3d). Stimulated PBMC from B→Orecipients showed a similar pattern of deficient B-specific IgMantibody (Table 1). IgG isotype antibodies against donor A and B

NATURE MEDICINE VOLUME 10 | NUMBER 11 | NOVEMBER 2004 1229

Table 1 Characteristics of ABO-incompatible and ABO-compatible heart transplant recipients

Pt. Age at time Current Blood Serum antibody Intragraft deposition Intragraft A- B- α-gal- A- B-No. of transplant age groups titer at time of of: expression of: specific specific specific specific specific

(years)a (donor→ transplant ELISA ELISA ELISA ELISPOT ELISPOTrecipient) A- B- C3 C4d IgG IgM A B

specific specific antigen antigen

1 2 mo 5.5 B→O 0 0 – – – – – + + – ND + –

2 7 wk 3.2 B→O 8 0 – – – – ND ND + – ND + –

3 2 d 5.6 B→O 0 0 – – – – – + + – ND + –

4 5 mo 3.0 A2B→O 128b 32b – – – – ND ND + – ND + –

5 1 mo 2.7 A→O 0 2 – – – – ND ND – ND + – ND

6 2 mo 6.6 A→O 0 0 – – – – + – – + ND – +

7 10 wk 4.8 A→O 1 1 – – – – + – – + + – +

8 5 mo 2.5c A→O 0 0 – – – – ND ND ND ND ND ND ND

9 14 mo 6.3 A→O 128 16 – – – – + – – + ND – +

10 25 d 8.4 AB→O 0 0 – – – – + + – – ND – –

11 2 mo 6.8 B→A 0 0 – – – – – + – – ND – –

12 2 mo 2.6 A→B 1 2 – – – – ND ND – – ND – –

13 8 mo 2.2 B→A 0 0 – – – – ND ND – – ND – –

14 5 mo 5.3 O→O —d —d ND ND ND ND ND ND + + ND + +

15 5 wk 4.7 O→O —d —d ND ND ND ND – – + + + + +

16 6 mo 8.4 B→O, O→Oe 2 0 ND ND ND ND – – + + ND + +

Median age at time of transplantation was 10.4 weeks for recipients of ABO-incompatible heart grafts (patients 1–13) and 20.4 weeks for recipients of ABO-compatible heartgrafts (patients 14–16). Median current age is 5.2 years for recipients of ABO-incompatible heart grafts and 5.3 years for recipients of ABO-compatible heart grafts. Interval sincetransplant ranges from 1.5 to 8.4 years for the group. Explanation of symbols: ND = test not done; + = positive; – = negative; mo, month; wk, week; d, day. aCurrent age as of June 2004. bPatient 4 was on ECMO support and receiving group O blood and plasma products. cPatient 8 expired at age 2.5 years due to accelerated graft vasculopathy.dSerum titers for patients 14 and 15 were presumed low because of age. ePatient 16 received a graft from a group O donor 3 d after receiving a graft from a group B donor.

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

antigens were absent for all patients tested (data not shown).These data suggest that donor-specific antibody deficiency inABO-incompatible recipients does not result from functionalinactivation of antigen-specific B cells.

Visualization of antibody-producing cellsTo detect B cells capable of producing donor-specific antibodies,we cultured patients’ PBMC with IL-2 and IL-4 stimulation. We visualized A-specific and B-specific IgM-producing cells withELISPOT assays (Table 1 and Fig. 4). We clearly observed spot formation showing B-specific but not A-specific antibody-producingcells in PBMC from an A→O recipients (Table 1 and Fig. 4a,b)whereas we visualized A-specific but not B-specific antibody-producingcells in PBMC from B→O and A2B→O recipients (Table 1 and Fig.4c,d). Both A-specific and B-specific antibody-producing cells wereobserved in PBMC from O→O recipients (Table 1 and Fig. 4e,f),although neither was visualized in PBMC from A→B, B→A andA1B→Ο recipients (Table 1). ELISPOT assays for IgG isotype antibody-producing cells were negative for all patients tested (datanot shown).

Detection of antigen-specific B lymphocytesThese studies suggest that B-cell elimination, not anergy, is the mech-anism of abrogation of B-cell responsiveness to donor A and B anti-gens in ABO-incompatible graft recipients. To directly ascertain thepresence of donor-specific B cells, we sought B lymphocytes bearingA antigen–specific B-cell receptors (BCR) by immunostainingpatients’ PBMC with biotin-conjugated synthetic group A antigenand FITC-conjugated human CD19-specific monoclonal antibody(Fig. 5). FACS analysis of PBMC from an O→O recipient showed1.8% of the total B-cell population with BCR specific for A antigen(Fig. 5a), whereas in PBMC from an A→O recipient, this B-cell popu-lation was absent (Fig. 5b), similar to the FACS profile of PBMC froma group A negative control (Fig. 5c).

DISCUSSIONThese studies of immunologic development, performed up to 8 yearsafter ABO-incompatible infant heart transplantation, show a persist-ent and selective deficiency of circulating antibodies to donor A andB antigens without antibody-mediated damage. Potential explana-tions for antibody deficiency include: intragraft immunoglobulindeposition (‘accommodation’), suppressed antibody productionresulting from nonspecific systemic immunosuppression and defi-cient antibody production resulting from specific B-cell tolerance.Absence of immunoglobulin and complement deposition in biopsyspecimens suggests that accommodation has not occurred; normaldevelopment of A-specific and B-specific antibodies in O→O infanttransplant recipients indicates that immunosuppressants do notsuppress isohemagglutinin development. Thus, the preponderanceof evidence suggests that these children have acquired donor-specificB-cell tolerance.

1230 VOLUME 10 | NUMBER 11 | NOVEMBER 2004 NATURE MEDICINE

10

A-s

peci

fic Ig

M (

ng/m

l)

A→O O→OA→O AControl

A→O O→OA→O AControl

20304050

6070

0

10

A-s

peci

fic Ig

M (

ng/m

l)

20304050

6070

0

A→O O→OA→O AControl

10

α-ga

l-spe

cific

IgM

(ng

/ml)

20304050

6070

0A→O O→OA→O A

Control

10

A-s

peci

fic Ig

M (

ng/m

l)

20304050

6070

0

a b

c dFigure 3 ELISA analyses of antibody in PBMC culture supernatantsshowing absence of A-specific antibody production by PBMC from two A→O recipients. (a) A-specific IgM isotype antibodies produced byPBMC cultured without stimulation. (b) A-specific IgM isotype antibodiesproduced by PBMC stimulated with IL-2 and IL-4 cytokines. (c) A-specificIgM isotype antibodies produced by PBMC stimulated with group Aerythrocytes. (d) Antibodies against ‘third-party’ carbohydrate epitope α-gal produced by IL-2- and IL-4-stimulated PBMC.

a b

c d

e f

A→O, A antigen A→O, B antigen

B→O, A antigen B→O, B antigen

Figure 2 Immunohistochemical staining showing absence of complementdeposition and persistence of donor antigens within endomyocardial biopsyspecimens from ABO-incompatible graft recipients. (a) Fluorescencestaining for complement component C4d in a specimen from an ABO-incompatible graft. (b) Similar staining for C4d in a graft specimen from aknown case of antibody-mediated rejection resulting from HLA sensitization(positive control). (c) A-antigen expression in a graft biopsy from a group Adonor 2 years after transplantation into a group O recipient. Arrows indicatepositive A-antigen staining. (d) Staining for B antigen in same patient as in c. (e) Staining for A antigen in a graft biopsy from a group B donor 4years after transplantation into a group O recipient. (f) B-antigen expressionin same patient as in e. Arrows indicate positive B-antigen staining.

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

Several B-cell tolerance mechanisms have been identified, includ-ing anergy (functional inactivation)29,30, deletion35–37 and receptorediting38–40. In vitro analysis of PBMC from ABO-incompatible graftrecipients showed that these cells were simultaneously incapable ofproducing antibodies against donor-type A and B antigens, yet capa-ble of producing antibodies against an irrelevant third-party polysac-charide antigen, and that B lymphocytes bearing BCR specific fordonor-type A and B antigens were absent. These data suggest thatanergy does not have a role in this setting and that antigen-specific Bcells have been eliminated by deletion or receptor editing.

The site at which the tolerizing interaction occurs between immature recipient B cells and donor A and B antigens is difficult todelineate in humans, thus definitive conclusions cannot be madefrom these experiments. Tolerance may be induced within the graft;alternatively, or in combination, tolerizing interactions may occur innongraft sites such as spleen or other lymphoid organs, or in bonemarrow or peripheral blood, through migration of graft-derived cellsor dispersal of donor antigens through perioperative administrationof donor-type noncellular blood products (platelets and plasma).Several findings suggest that transient exposure to donor antigens,such as administration of noncellular blood products, is insufficientto induce tolerance. First, patient 16 (blood group O) received a group B graft for a 3-d period only and was administered group B blood products intraoperatively. Yet after receiving a group O second graft, patient 16 developed B-specific antibody production to‘normal’ titers for age (Fig. 1h). Thus, transient exposure to B antigen of the first donor, whether in the graft itself or by group B blood products, was insufficient to induce tolerance. Second, patient 4(A2B→O) received group A1B platelets and plasma products contain-ing both A and B antigens. Yet, after transplantation with a group A2B graft, patient 4 was not rendered tolerant to A antigen containedin group A1B blood products, developing A-specific antibodies withnormal kinetics (Fig. 1a).

The spleen is thought to be important for A-specific and B-specific antibody production, inasmuch as splenectomy has beenreported to diminish the production of these antibodies in ABO-incompatible adult kidney transplant recipients1. But it has alsobeen shown that splenectomy in children does not prevent develop-ment of polysaccharide-specific antibodies41, showing that produc-tion of these antibodies is not limited to the spleen. In our one case ofa congenitally asplenic person (patient 6, A→O), B-specific antibodydeveloped normally (Fig. 1d), consistent with the theory that isohemagglutinin production is not limited to the spleen, whereas A-specific antibody production did not develop at all (Fig. 1c),suggesting that interactions leading to tolerance also are not confinedto the spleen.

These combined findings suggest, in accordance with animal toler-ance models27,28, that persistent exposure to donor antigens isrequired for tolerance induction, possibly in the graft itself, and thattolerance is dependent on a certain degree of antigen expression.Consistent with these results, experiments with 3-83 (H-2k,b-specific)immunoglobulin transgenic mice showed that specific self-reactive B cells are eliminated by deletion or receptor editing upon repeatedexposure to antigen-bearing cells42.

The developmental limits of susceptibility to tolerance induction to donor A and B antigens have not yet been delineated. Although

patient 9 (A→O) already had established A-specific antibody production prior to transplant at age 14 months (titer 1:128;Fig. 1c), A-specific antibody was depleted by plasma exchange during the operative procedure and did not reaccumulate to clinicallysignificant levels by 5 years after transplant, showing a persistent antibody deficiency similar to younger A→O recipients.Furthermore, A-specific antibody-producing cells could not bedetected by ELISPOT assay (Table 1). Thus, acquisition of ‘immunecompetence,’ shown by antibody production before transplant, didnot preclude continued susceptibility to apparent tolerance induc-tion. This raises the possibility that the window during which ABO-incompatible transplantation could be performed might beprolonged by repeated intentional exposure to nonself A and B antigens before transplant. This single case notwithstanding, cautionmust be advised for consideration of ABO-incompatible transplanta-tion in children in whom isohemagglutinin production has alreadybegun to develop.

Both systemic immunosuppression and (neonatal) thymectomyhave been shown to interfere with tolerance induction in experimen-tal settings43–47. That tolerance developed in ABO-incompatible

NATURE MEDICINE VOLUME 10 | NUMBER 11 | NOVEMBER 2004 1231

A→O, A-specific A→O, B-specific

B→O, A-specific B→O, B-specific

O→O, A-specific O→O, B-specific

a b

c d

e f

Figure 4 Visualization of antibody-producing cells by ELISPOT assayshowing absence of A-specific and B-specific antibody-producing cells incultured PBMC isolated from A→O (a,b), B→O (c,d) and O→O (e,f) hearttransplant recipients. Spots indicate specific antibody-producing cells.

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

infant recipients despite these factors may be related to the fact that polysaccharide antigens such as blood group substances are considered to be ‘T-cell independent’48, and most systemicimmunosuppressive drugs are directed predominantly at T-cell function, which is largely mature in neonates49.

Whether tolerance to donor A and B antigens in infants is alsoassociated with tolerance to donor HLA has not been addressed inthese studies. Although tolerance may develop to donor HLA,one could also speculate that the developmental differences in maturation of human immune responses to carbohydrates versusproteins19,21 may result in susceptibility to tolerance induction tocarbohydrate antigens (A and B) concomitant with ‘sensitization’to protein antigens (HLA).

Neonatally induced transplantation tolerance has not beenshown previously in humans. The setting of ABO-incompatibleinfant heart transplantation is unique in that it allows an earlyimmune intervention and a clear strategy of assessing immunityversus tolerance to defined donor antigen(s). We have delineatedthe mechanism of successful ABO-incompatible infant hearttransplantation as “actively acquired tolerance of foreign cells”9,and characterized here as donor antigen–specific B-cell elimina-tion. For the first time, we have shown that human infants are suf-ficiently immature for intentional induction of immunologictolerance, at least in the B-cell compartment to T-independent A and B antigens. Through understanding the mechanisms under-lying tolerance in this setting, it may be possible to expand thepotential applications of this strategy to older patient populations,which could result in decreased mortality for patients awaitingtransplantation and increased usage of donor organs.Additionally, our findings have significance for xenogeneic trans-plantation in which similar issues of polysaccharide antigen andantibody production remain major obstacles to success50.

METHODSPatients and biologic samples. We studied 16 cardiac allograft recipientsbetween February 1996 and October 2002 (Table 1); 13 infants (median ageat transplant 10.4 weeks) received grafts from ABO-incompatible donorsand 3 infants (median age 20.4 weeks) received ABO-compatible grafts. Ofthe 16 transplant recipients, 15 recipients are clinically well at current agesranging from 2.2 to 8.4 years; one recipient died (age 2.5 years) due to accel-erated graft vasculopathy (undetectable donor-specific antibody at time ofdeath). We reported heart transplant and immunosuppression details previ-ously24. Near-total thymectomy was performed as a routine part of infantcardiac surgery; splenectomy was not performed. We obtained peripheralserum samples for isohemagglutinin detection at defined time points after transplant. Endomyocardial biopsies for clinical rejection surveillance were

obtained at defined time points; we performed immunohistochemistrystaining to detect antibody-mediated rejection and donor antigen persist-ence. For a subset of 15 transplant recipients (Table 1), additional wholeblood samples were obtained for B-cell functional analysis. We performedexperiments in compliance with institutional guidelines and approved bythe Research Ethics Board of The Hospital for Sick Children. Informed con-sent was obtained from transplant recipients’ parents.

Blood group determination and isohemagglutinin assay. We determined bloodtypes using standard blood bank procedures. Serum A-specific and B-specificantibody titers were measured using agglutination tests with erythrocytes ofknown blood type (‘reverse’ blood typing). We diluted patient serum with salinein ratios ranging from 1:1 to 1:256 and mixed the serum with group A or B erythrocytes. A modified version of the isohemagglutination assay was used totest for serum factors that might inhibit erythrocyte agglutination by A-specificor B-specific antibodies (Supplementary Methods online).

Immunohistochemical staining. We stained endomyocardial biopsy sectionsfor deposition of immunoglobulin and complement components and for persistent expression of donor A or B antigens (Supplementary Methodsonline) and a cardiac immunopathologist examined the sections.

Cell isolation and cultures. We layered patient peripheral blood ontoHISTOPAQUE-1077 solution (Sigma). Following centrifugation, we removedserum and stored it at –30 °C. PBMC at the serum-HISTOPAQUE interfacewere washed and either used immediately for cell culture or flow cytometry, orresuspended in fetal bovine serum (FBS) containing 10% dimethyl sulfoxide(Sigma) for long-term storage in liquid nitrogen. We cultured 1 × 105 cells perwell in RPMI 1640 medium supplemented with 10% FBS, 5 µg/ml bovineinsulin (Sigma), 5 µg/ml human transferrin (Sigma), 5 ng/ml sodium selenite,50 µM 2-mercaptoethanol, 100 U/ml penicillin and 100 µg/ml streptomycin.For erythrocyte-stimulated cultures, 2 × 105 cells per well of washed humanerythrocytes were added on the first day of culture. We did not changemedium before supernatant collection. In cytokine-stimulated cultures, 200U/ml human recombinant interleukin(IL)-2 (PeproTech Canada) and 50U/ml human recombinant IL-4 (PeproTech Canada) were added on the firstday of culture.

Enzyme-linked immunosorbent assay (ELISA). We coated polystyreneplates (Corning) with 5 µg/ml synthetic group A or B trisaccharide conju-gated to bovine serum albumin (V-Labs). Nonspecific binding sites wereblocked with PBS-Tween containing 2% FBS. We cultured patients’cryopreserved PBMC with or without stimulation. After 10 d, culture supernatants were incubated in wells for 1 h; bound antibodies weredetected using horseradish peroxidase–conjugated goat antibodies tohuman IgM or IgG (Southern Biotechnology, 1:4,000 dilution) followed bycolor development with 3,3’,5,5′-tetramethylbenzidine (Sigma). Westopped the reaction with sulfuric acid after 30 min and measuredabsorbance at 450 nm. Preparation of a standard curve allowed antibodyconcentration to be determined from optical density. Negative controlswere PBMC from blood group A (A-specific negative control) and bloodgroup B (B-specific negative control) volunteers; assay results for negativecontrol samples were 0–4 ng/ml; assay results >5 ng/ml were scored as positive.

Enzyme-linked immunospot (ELISPOT) assay. We coated nitrocellulose mem-branes of 96-well MultiScreen-HA plates (Millipore) with 5 µg/ml synthetic group A or B trisaccharide conjugated to bovine serum albumin.Cryopreserved PBMC were cultured for 10 d with IL-2 and IL-4. After washing,we cultured cells for 24 h in MultiScreen-HA plates and stained them with biotin-conjugated goat antibodies to human IgM or IgG (Southern Biotechnology,1:4,000 dilution). Incubation with streptavidin-horseradish peroxidase was fol-lowed by color development with AEC solution. Negative controls, in which wedid not visualize any spots, were PBMC from blood group A (A-specific negativecontrol) and blood group B (B-specific negative control) volunteers.

Flow cytometry. We immunostained PBMC for direct detection of B lymphocytes bearing A antigen–specific BCR. PBMC (5 × 105 cells) were incubated with 1.25 µg/ml synthetic group A tetrasaccharide-PAA-biotin

1232 VOLUME 10 | NUMBER 11 | NOVEMBER 2004 NATURE MEDICINE

Group A antigen-biotin/streptavidin-PE

CD

19-F

ITC

1.8% <0.2% <0.2%

100

101

102

103

104

100 101 102 103 104

100

101

102

103

104

100

101

102

103

104

100 101 102 103 104 100 101 102 103 104

a b c

Figure 5 Detection by flow cytometry of B lymphocytes bearing BCRspecific to A antigen in PBMC isolated from infant heart transplantrecipients. Each graph contains 104 gated CD19+ cells. Percentagesindicate the proportion of the CD19+ population bearing A antigen–specificBCR. Panels show analyses of PBMC from (a) an O→O recipient (positivecontrol patient) (b) an A→O recipient and (c) a group A negative control.FITC, fluorescein isothiocyanate; PE, phyloerythrin.

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine

A R T I C L E S

conjugate (GlycoTech) and FITC-conjugated human CD19-specific monoclonal antibody (PharMingen, 10 µl/106 cells), followed by strep-tavidin-phycoerythrin. We used group A and O PBMC as negative and positive controls, respectively, with at least two samples for each experimentalgroup. We gated CD19+ cells using FACScan flow cytometer (BD Biosciences)and analyzed data with WinMDI Version 2.8. The detection limit of group A antigen-specific BCR-bearing cells in the negative control population wasapproximately 0.2% of total CD19+ B cells.

Note: Supplementary information is available on the Nature Medicine website.

ACKNOWLEDGMENTSWe thank K. Wood and M. Sykes for discussion and advice, and P. Morris and R.Zhong for reading of our manuscript. This work was supported with funding fromthe Canadian Institutes for Health Research, the Heart and Stroke Foundation ofOntario, and the Research Training Competition at The Hospital for Sick Children.

COMPETING INTERESTS STATEMENTThe authors declare that they have no competing financial interests.

Received 3 June; accepted 1 October 2004Published online at http://www.nature.com/naturemedicine/

1. Alexandre, G.P. et al. Present experiences in a series of 26 ABO-incompatible livingdonor renal allografts. Transplant. Proc. 19, 4538–4542 (1987).

2. Sutherland, D.E. et al. Long-term effect of splenectomy versus no splenectomy inrenal transplant patients. Reanalysis of a randomized prospective study.Transplantation 38, 619–624 (1984).

3. Bannett, A.D., McAlack, R.F., Morris, M., Chopek, M.W. & Platt, J.L. ABO incom-patible renal transplantation: a qualitative analysis of native endothelial tissue ABOantigens after transplantation. Transplant. Proc. 21, 783–785 (1989).

4. Chopek, M.W., Simmons, R.L. & Platt, J.L. ABO-incompatible kidney transplanta-tion: initial immunopathologic evaluation. Transplant. Proc. 19, 4553–4557(1987).

5. Cooper, D.K. A clinical survey of cardiac transplantation between ABO blood group-incompatible recipients and donors. Transplant Proc. 22, 1457 (1990).

6. Gugenheim, J., Samuel, D., Reynes, M. & Bismuth, H. Liver transplantation acrossABO blood group barriers. Lancet 336, 519–523 (1990).

7. Owen, R.D. Immunogenetic consequences of vascular anastomoses betweenbovine twins. Science 102, 400–405 (1945).

8. Burnet, F.M. & Fenner, F. The Production of Antibodies. (Macmillan and Company,Melbourne, 1949).

9. Billingham, R.E., Brent, L. & Medawar, P.B. Activity acquired tolerance of foreigncells. Nature 172, 603–606 (1953).

10. Dorsch, S. & Roser, B. Suppressor cells in transplantation tolerance. I. Analysis ofthe suppressor status of neonatally and adoptively tolerized rats. Transplantation33, 518–524 (1982).

11. Kappler, J.W., Roehm, N. & Marrack, P. T cell tolerance by clonal elimination in thethymus. Cell 49, 273–280 (1987).

12. Klinman, N.R. The “clonal selection hypothesis” and current concepts of B cell tol-erance. Immunity 5, 189–195 (1996).

13. McCarthy, S.A. & Bach, F.H. The cellular mechanism of maintenance of neonatallyinduced tolerance to H-2 class I antigens. J. Immunol. 131, 1676–1682 (1983).

14. Ramsdell, F. & Fowlkes, B.J. Clonal deletion versus clonal anergy: the role of thethymus in inducing self tolerance. Science 248, 1342–1348 (1990).

15. Ramsdell, F., Lantz, T. & Fowlkes, B.J. A nondeletional mechanism of thymic selftolerance. Science 246, 1038–1041 (1989).

16. Streilein, J.W. Neonatal tolerance of H-2 alloantigens. Procuring graft acceptancethe “old-fashioned” way. Transplantation 52, 1–10 (1991).

17. West, L.J. Defining critical windows in the development of the human immune sys-tem. Hum. Exp. Toxicol. 21, 499–505 (2002).

18. West, L.J. Developmental aspects of immunomodulation: exploiting the immatureimmune system for organ transplantation. Transpl. Immunol. 9, 149–153 (2002).

19. Cadoz, M. Potential and limitations of polysaccharide vaccines in infancy. Vaccine16, 1391–1395 (1998).

20. Gennery, A.R. et al. Effect of immunosuppression after cardiac transplantation inearly childhood on antibody response to polysaccharide antigen. Lancet 351,1778–1781 (1998).

21. Wilson, C.B. Immunologic basis for increased susceptibility of the neonate to infec-tion. J. Pediatr. 108, 1–12 (1986).

22. Fong, S.W., Qaqundah, B.Y. & Taylor, W.F. Developmental patterns of ABO isoag-

glutinins in normal children correlated with the effects of age, sex., and maternalisoagglutinins. Transfusion 14, 551–559 (1974).

23. Springer, G.F. & Horton, R.E. Blood group isoantibody stimulation in man by feed-ing blood group-active bacteria. J. Clin. Invest. 48, 1280–1291 (1969).

24. West, L.J. et al. ABO-incompatible heart transplantation in infants. N. Engl. J.Med. 344, 793–800 (2001).

25. Smalley, C.E. & Tucker, E.M. Blood group A antigen site distribution andimmunoglobulin binding in relation to red cell age. Br. J. Haematol. 54, 209–219(1983).

26. Mollison, P.L., Contreras, M. & Engelfriet, C.P. ABO, Lewis, Ii and P groups. inBlood Transfusion in Clinical Medicine. 9th edn 155 (Blackwell ScientificPublishing, Oxford, 1993).

27. Hamano, K., Rawsthorne, M.A., Bushell, A.R., Morris, P.J. & Wood, K.J. Evidencethat the continued presence of the organ graft and not peripheral donormicrochimerism is essential for maintenance of tolerance to alloantigen in vivo inanti-CD4 treated recipients. Transplantation 62, 856–860 (1996).

28. Morecki, S., Leshem, B., Eid, A. & Slavin, S. Alloantigen persistence in inductionand maintenance of transplantation tolerance. J. Exp. Med. 165, 1468–1480(1987).

29. Erikson, J., Radic, M.Z., Camper, SA., Hardy, R.R., Carmack, C. & Weigert, M.Expression of anti-DNA immunoglobulin transgenes in non-autoimmune mice.Nature 349, 331–334 (1991).

30. Goodnow, C.C. et al. Altered immunoglobulin expression and functional silencingof self-reactive B lymphocytes in transgenic mice. Nature 334, 676–682 (1988).

31. Chelvarajan, R.L., Gilbert, N.L. & Bondada, S. Neonatal murine B lymphocytesrespond to polysaccharide antigens in the presence of IL-1 and IL-6. J. Immunol.161, 3315–3324 (1998).

32. Luo, W., Fine, J., Garg, M., Kaplan, A.M. & Bondada, S. Interleukin-10 enhancesimmune responses to pneumococcal polysaccharides and sheep erythrocytes inyoung and aged mice. Cell Immunol. 195, 1–9 (1999).

33. Splawski, J.B. & Lipsky, P.E. Cytokine regulation of immunoglobulin secretion byneonatal lymphocytes. J. Clin. Invest. 88, 967–977 (1991).

34. Galili, U. Interaction of the natural anti-Gal antibody with alpha-galactosyl epi-topes: a major obstacle for xenotransplantation in humans. Immunol. Today 14,480–482 (1993).

35. Chen, C. et al. The site and stage of anti-DNA B-cell deletion. Nature 373,252–255 (1995).

36. Hartley, S.B., Crosbie, J., Brink, R., Kantor, A.B., Basten, A. & Goodnow, C.C.Elimination from peripheral lymphoid tissues of self-reactive B lymphocytes recog-nizing membrane-bound antigens. Nature 353, 765–769 (1991).

37. Nemazee, D.A. & Burki, K. Clonal deletion of B lymphocytes in a transgenic mousebearing anti-MHC class I antibody genes. Nature 337, 562–566 (1989).

38. Gay, D., Saunders, T., Camper, S. & Weigert, M. Receptor editing: an approach byautoreactive B cells to escape tolerance. J. Exp. Med. 177, 999–1008 (1993).

39. Radic, M.Z., Erikson, J., Litwin, S. & Weigert, M. B lymphocytes may escape toler-ance by revising their antigen receptors. J. Exp. Med. 177, 1165–1173 (1993).

40. Tiegs, S.L., Russell, D.M. & Nemazee, D. Receptor editing in self-reactive bonemarrow B cells. J. Exp. Med. 177, 1009–1020 (1993).

41. Hammarstrom, L. & Smith, CI. Development of anti-polysaccharide antibodies inasplenic children. Clin. Exp. Immunol. 66, 457–462 (1986).

42. Lang, J. & Nemazee, D. B cell clonal elimination induced by membrane-boundself-antigen may require repeated antigen encounter or cell competition. Eur. J.Immunol. 30, 689–696 (2000).

43. Wang, C., Sun, J., Sheil, A.G., McCaughan, G.W. & Bishop, G.A. A short course ofmethylprednisolone immunosuppression inhibits both rejection and spontaneousacceptance of rat liver allografts. Transplantation 72, 44–51 (2001).

44. Perico, N., Imberti, O., Bontempelli, M. & Remuzzi, G. Immunosuppressive ther-apy abrogates unresponsiveness to renal allograft induced by thymic recognition ofdonor antigens. J. Am. Soc. Nephrol. 5, 1618–1623 (1995).

45. Yamada, K. et al. Role of the thymus in transplantation tolerance in miniatureswine. I. Requirement of the thymus for rapid and stable induction of tolerance toclass I-mismatched renal allografts. J. Exp. Med. 186, 497–506 (1997).

46. Noris, M. et al. Thymic microchimerism correlates with the outcome of tolerance-inducing protocols for solid organ transplantation. J. Am. Soc. Nephrol. 12,2815–2826 (2001).

47. Nakafusa, Y., Goss, J.A. & Flye, M.W. Prevention by thymectomy of toleranceinduced by intrathymic injection of donor splenocytes. Surgery 114, 183–189(1993).

48. Mond, J.J., Lees, A. & Snapper, C.M. T cell-independent antigens type 2. Annu.Rev. Immunol. 13, 655–692 (1995).

49. Holsapple, M.P., West, L.J. & Landreth, K.S. Species comparison of anatomicaland functional immune system development. Birth Defects Res. Part B Dev.Reprod. Toxicol. 68, 321–334 (2003).

50. Lin, S.S. et al. The role of anti-Galalpha1-3Gal antibodies in acute vascular rejec-tion and accommodation of xenografts. Transplantation 70, 1667–1674 (2000).

NATURE MEDICINE VOLUME 10 | NUMBER 11 | NOVEMBER 2004 1233

©20

04 N

atur

e P

ublis

hing

Gro

up

http

://w

ww

.nat

ure.

com

/nat

urem

edic

ine