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. 2002 Mar;127(3):412–422. doi: 10.1046/j.1365-2249.2002.01789.x

Feeding NOD mice with pig splenocytes induces transferable mechanisms that modulate cellular and humoral xenogeneic reactions against pig spleen or islet cells

S YOU 1, E GOUIN 1, P SAÏ 1
PMCID: PMC1906308  PMID: 11966756

Abstract

We have reported previously that oral administration of pig cells to NOD mice modified xenogeneic cellular response against pig islet cells (PICs), and hypothesized that it may have induced active suppression. This preliminary report evaluated only the effect of feeding pig cells by ‘primary’ proliferation, i.e. when splenocytes from fed mice are confronted with pig cells in vitro. The present study also considered ‘secondary’ proliferation and cytokine production after feeding and subsequent in vivo graft of pig cells. Additionally, serum IgM and IgG isotypes were quantified by ELISA using pig target cells. Induction of active mechanism by feeding was hypothetical, which led us here to transfer splenocytes from mice fed pig spleen cells (PSC) and evaluate ‘primary’ (after transfer) and ‘secondary’ (after transfer and subsequent graft of pig cells) proliferations and cytokine secretions in recipient mice. We also determined whether the effects of feeding pig cells persisted after depression of suppressor mechanisms by cyclophosphamide. Mice fed with PSC displayed increased ‘primary’ splenocyte proliferation to PSC or PIC (P < 0·0001), while ‘secondary’ responses were decreased (P < 0·03) in those fed PSC and subsequently grafted with PSC. The increased ‘primary’ and decreased ‘secondary’ proliferations were reduced (P < 0·04) by pretreatment with cyclophosphamide. The IL-10/ and IL-4/IFNγ ratios produced in response to PSC increased (P < 0·04) in mice fed and grafted with PSC compared to those grafted only with PSC. IgM and IgG levels against pig cells were, respectively, increased (P < 0·04) and decreased (P < 0·04) in mice fed and grafted with PSC. IgG2a and IgG2b, but not IgG1, levels were lower (P < 0·01). These effects of feeding PSC on ‘secondary’ proliferation, cytokine and antibody productions, were not detected when mice were fed PSC only after graft with PSC. Transfer with splenocytes from mice fed PSC increased ‘primary’ proliferation of splenocytes from recipient mice in response to PSC (P < 0·02) or PIC (P < 0·05). After transfer with splenocytes from PSC-fed mice and graft with PSC, ‘secondary’ proliferation to pig cells were reduced (P < 0·04), and the IL-10/IFNγ ratio produced in response to PSC was increased fourfold. Thus, oral administration of PSC induces active transferable mechanisms, characterized by a biphasic pattern with early increased ‘primary’ xenogeneic cellular reactions to both PSC and PIC, followed by decreased ‘secondary’ responsiveness and a concomitant shift of the Th1/Th2 balance towards greater Th2 influence. Decreased responsiveness may be due to active suppression, even though induction of anergy or deletion cannot be excluded.

Keywords: cellular reaction, cyclophosphamide, humoral response, islets, mouse, oral administration, pig, spleen cells, transfer, xenograft

INTRODUCTION

We have reported previously [1] that oral administration of pig cells to non-obese diabetic (NOD) mice, a model for type 1 diabetes [2], constitutes a novel approach for modification of xenogeneic cellular response to pig islet cells (PIC) [3], and more generally that feeding of cells from xenogeneic species is not without consequences to the immune system. Mice fed with pig spleen cells (PSC) displayed increased proliferation of splenocytes in response to PSC or PIC. Concomitantly, splenocytes from control mice inhibited basal and stimulated insulin releases from PIC in vitro, which is indicative of aggression, whereas this inhibition was reduced during coincubation with splenocytes from mice fed with PSC. By analogy with the down-regulation of systemic immune responses by oral administration of autoantigens [4–7], particularly in the NOD mouse [8–10], and allogeneic cells [11–15] we thus suggested that feeding NOD mice with PSC may have been conducive to active suppression. However, this preliminary study lacked several types of data needed to improve our understanding of the mechanisms involved during oral administration of pig cells, and especially during subsequent systemic contact with pig cells, as would occur after a xenograft.

First, the effect of oral administration of PSC on cellular xenogeneic reaction was evaluated only by ‘primary’ proliferation, i.e. when splenocytes from fed mice were confronted for the first time with pig cells in vitro. In the present work, ‘secondary’ splenocyte proliferation and cytokine production in response to pig cells were also studied in mice fed and subsequently grafted with PSC. Additionally, as our previous report was concerned only with cellular immunity, systemic antibody response resulting from the contact of the digestive immune system with PSC was not studied. Thus, an ELISA assay was designed in the present study, using pig cells as targets, to quantify IgM and IgG isotypes after feeding PSC. Analyses of IgG isotypes and cytokine production were also designed as approaches to the modification of the Th2/Th1 balance induced by feeding PSC. Furthermore, as the active cellular mechanisms induced through the feeding of PSC were only hypothetical, it was necessary to determine whether these mechanisms actually existed. Thus, after transfer of splenocytes from mice fed PSC, ‘primary’ (after transfer alone) and ‘secondary’ (after transfer and subsequent graft of PSC) proliferations and cytokine productions in response to pig cells were evaluated in recipient mice. Finally, as another means of clarifying whether suppressor cellular mechanisms were induced, we investigated whether the effects of feeding PSC persisted after depression of suppressor mechanisms by cyclophosphamide [16].

MATERIALS AND METHODS

Specific pathogen-free (SPF) pigs: islet and splenocyte isolations

Pancreata were removed from Large White SPF pigs (80–120 kg, 20–30 weeks old), soaked in betadine solution, inflated immediately with modified University of Wisconsin (mUW) solution, transported in a receptacle containing mUW, inflated again with mUW solution containing 2 mg/ml liberase (Roche Diagnostics, Meylan, France), and placed in a digestion chamber filled with mUW solution kept at 36°C [17–20]. Crude islets were pelleted by centrifugation, suspended in mUW and purified on continuous Optiprep gradient (Life International Technology, Cergy-Pontoise, France) with a COBE 2991 processor. Islet purity, as assessed by dithizone staining, exceeded 90%. Insulin release from SPF pig islets in response to different stimuli was highly reproducible. Islets were treated with EDTA and dispase to obtain islet cells suspended in Ham’s F10 medium (5·5 mmol/l glucose) (Biomédia, Boussens, France) supplemented with 2% Ultroser (BioSepra SA, Villeneuve la Garenne, France), penicillin, streptomycin and Fungizone (Seromed, Berlin, Germany). Pig spleens were removed in Hanks’s balanced salt solution (HBSS) (Seromed), splenocytes were flushed out and suspended in RPMI (Eurobio, Les Ulis, France).

Feeding NOD mice with pig splenocytes

NOD mice (Clea, Tokyo, Japan) have been bred in our laboratory since 1986. Two-month-old non-diabetic female mice were used. Pig spleen cells (PSC), prepared as lysates by a single freezing and thawing cycle and suspended in phosphate-buffered saline (PBS), were administered orally in a volume of 0·25 ml per mouse per feeding. Daily oral administrations of 25 × 106 PSC or PBS were performed for 11 days. In some experiments, two intraperitoneal injections of cyclophosphamide (Cy, Endoxan-Asta™, Laboratoires Asta-Medica, Mérignac, France) were administered 24h prior to and 14 days after the first feeding, at doses (300 and 200 mg/kg) known to depress suppressor mechanisms in NOD mice [16]. At the end of the experiments, serum samples and spleens were removed. Splenocytes were flushed out and suspended in RPMI.

Effect of feeding PSC on ‘primary’ and ‘secondary’ proliferations of splenocytes

‘Primary’ proliferation of mouse splenocytes to PSC or PIC were tested 10 days after the last gavage. For analysis of ‘secondary’ proliferation, mice were fed PSC and exposed twice intraperitoneally to 15 × 106 PSC on days 6 and 16 after the first feeding. The first exposure was performed in complete Freund’s adjuvant (CFA), and the second in incomplete Freund’s adjuvant (IFA) (Sigma Immunochemicals, St Quentin Fallavier, France). Control mice were exposed to these vehicles. Under similar conditions to those described above, in vitro proliferations of mouse splenocytes to PSC or PIC were then tested 7 days after the second i.p. exposure. In some experiments, ‘secondary’ proliferations were analysed when mice were fed PSC only after i.p. exposure to PSC instead of prefeeding (feedings were began on day 4 after the second i.p. exposure; proliferations were tested 10 days after the last feeding). Spleen cells were aliquoted into flat-bottomed 96-well plates (NUNC, Roskilde, Denmark) at 0·5 × 106/well with 60 000 PSC or PIC in 0·3 ml RPMI containing 1·5% autologous decomplemented mouse serum. Plates were incubated for 6 days and pulsed by adding 3·7 × 104[3H]thymidine (sp. act. 7·4 × 108/mmol; Amersham, Les Ulis, France) for the final 18 h. Cells were harvested and counted. Data were obtained from the mean values of triplicate wells. The results were also expressed by a stimulation index (SI), i.e. the ratio between the count with stimulator pig cells and the basal count. The interassay coefficient of variation (s.d./mean × 100) of the SI ranged from 5 to 8%.

Effect of transfer of splenocytes from mice fed PSC on splenocyte proliferations of recipient mice

Splenocytes were isolated 1 day after the last feeding from fed mice and transferred three times (days 0, 3 and 13) i.v. to syngeneic recipients (70 × 106 cells/transfer). In some experiments, recipient mice were irradiated (750 R) 24h prior to the first transfer. To investigate the effect of transfer on ‘primary’ proliferation of splenocytes from recipient mice, responses to PSC or PIC were tested on day 23 after the first transfer. To evaluate the effect on ‘secondary’ proliferation, transferred recipient mice were then exposed twice i.p. to 15 × 106 PSC on days 6 and 16 after the first transfer, and proliferation of splenocytes was tested on day 23 after the first transfer.

‘Primary’ and ‘secondary’ cytokine secretion from mouse splenocytes in response to pig cells

Under similar conditions to those described above for proliferations, 100 μl of supernatant were removed from each well after 2 days of co-incubation for interferon-γ (INFγ), interleukin-10 (IL-10) and IL-4 assays. Cytokine concentrations were determined by ELISA (R&D Systems, Abingdon, UK). Standard curves were generated using recombinant mouse interleukins.

Effect of feeding PSC on humoral response to pig cells (ELISA)

PSC were suspended in PBS, pH 7·2 (0·4 × 106 cells/ml), and distributed (50 μl/well) into 96-well flat-bottomed plates (NUNC). Wells without cells provided a measure of nonspecific binding of reagents to the plastic. Plates were dried overnight at 37°C. After storage, 200 μl of PBS containing 1% bovine serum albumin and 0·3% gelatin (Sigma, St Louis, MO, USA) were added for 90 min. Diluted mouse sera (50 μl) were then added in duplicate for 2 h:1:50 to 1:200 for IgM and IgG2a, and 1:400 to 1:1600 for IgG, IgG1 and IgG2b. After four washes with PBS-0·1% BSA-0·3% gelatin, 50 μl of alkaline phosphatase-conjugated (antimouse IgM) or peroxidase-conjugated antibodies (anti-IgG, anti-IgG1, anti-IgG2a, anti-IgG2b) diluted in PBS-0·1% BSA were allowed to react for 2 h. The conjugated antibodies were used at the following dilutions: anti-IgM (1:1000) (Sigma), anti-IgG (1:5000) (Sigma), anti-IgG1 (1:2000), anti-IgG2a (1:1500) and anti-IgG2b (1:2000) (Tebu, Le Perray-en-Yveline, France). After four washes, 200 μl of p-nitrophenylphosphate or o-phenylenediamine dihydrochloride substrate solutions (Sigma) were added. All steps were performed at room temperature. The reaction was stopped after 30 min by addition of 3 N sodium hydroxide or chloridric acid. Optical densities (O.D.) at 405 nm or 492 nm were determined (Titertek Multiskan, Flow Laboratoiries, McLean, VA, USA). Results were also expressed as index values = O.D. [sample serum] − O.D. [negative standard serum]/O.D. positive standard serum] − O.D. [negative standard serum].

Stastistical analysis

Data are presented as mean values ± s.e.m. The statistical significance of differences was evaluated using Student’s t-test or Mann–Whitney U-test. P < 0·05 was considered significant.

RESULTS

Effects of feeding pig spleen cells (PSC) on ‘primary’ and ‘secondary’ proliferations of mouse splenocytes in response to pig cells

Compared to NOD mice fed PBS, those fed with PSC displayed much stronger (P < 0·0001) splenocyte ‘primary’ proliferation (Fig. 1a) in response to PSC (SI: 2·3 ± 0·3 and 11·8 ± 1·5, respectively) and PIC (SI: 4·3 ± 0·4 and 11·4 ± 1·3, respectively). A positive relationship was found between increased ‘primary’ splenocyte proliferation in response to either PSC (regression analysis, P < 0·001) or PIC cells (P < 0·02) and the number of cells per feeding (data not shown). Proliferation increased up to the highest number of cells per feeding (200 × 106), and no hyporesponsiveness could be detected with this very high number of cells per feeding.

Fig. 1.

Fig. 1

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Effects of feeding pig spleen cells (PSC) on ‘primary’ or ‘secondary’ proliferations of NOD mouse splenocytes in response to PSC or islet cells (PIC) in vitro. Daily administrations of 25 × 106 PSC or PBS were given for 11 days. For analysis of splenocyte ‘primary’ responsiveness to PSC or PIC (a), mice fed PBS (control feeding, n = 58) or PSC (n = 61) were studied 10 days after the last feeding. For analysis of ‘secondary’ proliferation (b), mice (n = 11 in each group) prefed PBS or PSC and grafted twice intraperitoneally with 15 × 106 PSC (or vehicles) on days 6 and 16 after the first feeding were studied 7 days after the second i.p. exposure. (c) The decrease of ‘secondary’ proliferation to PSC and PIC was not detected when mice (n = 3) were fed PSC only after i.p. graft with PSC (instead of the prefeeding shown in (b)). The results indicate the mean ± s.e.m. of [3H]thymidine uptake by 0·5 × 106 splenocytes alone (open bars) or co-incubated for 6 days with 60 000 pig cells (black bars). The mean ± s.e.m. of the stimulation index (SI = the ratio of the mean count with stimulator cells to the mean basal count) was determined. * P < 0·04, ** P < 0·03 and *** P < 0·0001 indicate significant differences between mice fed with PSC or PBS.

Mice fed and subsequently grafted with PSC displayed lower (P < 0·03) ‘secondary’ splenocyte proliferation in response to PSC than did those grafted with PSC but prefed PBS (Fig. 1b). The same mice also displayed decreased (P < 0·04) secondary splenocyte proliferation in response to PIC (Fig. 1b). The decreased secondary splenocyte proliferation observed in fed mice receiving two i.p. injections of PSC was also observed in fed mice given a single injection of PSC (data not shown). These effects of prefeeding PSC were durable as they were observed throughout the experiment (up to 10 weeks after the last feeding). The decrease in ‘secondary’ proliferation to PSC and PIC was not detected when mice were fed PSC only after graft with PSC instead of prefeeding (Fig. 1c).

Cyclophosphamide pretreatment of mice fed with PSC (Fig. 2) reduced the enhancing effect on primary splenocyte proliferation to PSC or PIC (P < 0·04) and the decreasing effect on ‘secondary’ proliferation to PSC (P < 0·01) or PIC (P < 0·04).

Fig. 2.

Fig. 2

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Influence of cyclophosphamide (Cy) treatment relative to the effects of feeding pig spleen cells (PSC) on ‘primary’ and ‘secondary’ splenocyte proliferations in response to PSC or pig islet cells (PIC) in vitro. NOD mice were treated or not with Cy (see Materials and methods) and fed PBS or PSC (25 × 106 PSC/day for 11 days). Splenocyte ‘primary’ responsiveness to PSC or PIC (a) was then analysed in some mice (n = 10 per group) 10 days after withdrawal of oral treatment. For analysis of ‘secondary’ response (b), other mice (n = 10 per group) were i.p.-grafted (15 × 106 PSC on days 6 and 16 after the first feeding), and splenocyte proliferations were tested 7 days after the second i.p. exposure. The results indicate the mean ± s.e.m. of [3H]thymidine uptake by 0·5 × 106 splenocytes alone (open bars) or coincubated for 6 days with pig cells (black bars). The mean ± s.e.m. of the stimulation index (SI = the ratio of the mean count with stimulator cells to the mean basal count) was determined. * P < 0·04 and ** P < 0·01 indicate significant differences between mice treated or not with Cy.

Effect of feeding PSC on ‘primary’ and ‘secondary’ cytokine secretion from mouse splenocytes in response to pig cells

During ‘primary’ proliferation in response to PSC, IL-10, IL-4 and IFNγ productions increased in PSC-fed mice compared to control mice (Fig. 3a). However, as IL-10 and IL-4 productions increased more than IFNγ production (5·8-fold and 4·9-fold versus 1·8-fold), the IL-10/IFNγ and IL-4/IFNγ ratios were increased.

Fig. 3.

Fig. 3

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Effect of feeding pig spleen cells (PSC) on ‘primary’ and ‘secondary’ cytokine secretions from NOD mouse splenocytes in response to PSC. Daily administrations of 25 × 106 PSC or PBS were given for 11 days. ‘Primary’ splenocyte IL-10, IL-4 and IFNγ secretions (a) were analysed 12 days after the last feeding (n = 3 in each group). ‘Secondary’ cytokine secretions (b) were analysed 7 days after the second i.p. exposure in mice (n = 3 in each group) prefed PBS or PSC and i.p.-grafted (15 × 106 PSC, or the vehicles, on days 6 and 16 after the first feeding). (c) The changes in ‘secondary’ cytokine secretions in response to pig cells were not detected when mice (n = 3) were fed PSC only after i.p. graft with PSC (instead of prefeeding shown in (b)). The results indicate the mean ± s.e.m. production of IFNγ and IL-10 by 0·5 × 106 splenocytes alone (□) or co-incubated for 48 h with 60 000 pig cells (■). The means ± s.e.m. of the IL-10/IFNγ and IL-4/IFNγ ratios are also indicated. * P < 0·04 between mice fed with PSC or PBS.

The main change in cytokine production was observed during ‘secondary’ proliferations (Fig. 3). In control mice fed PBS and subsequently grafted with PSC, IL-10, IL-4 and IFNγ productions increased in response to PSC; the increase of IFNγ (110-fold) was higher than that of IL-10 (14·5-fold) and IL-4 (10·1-fold). Compared to these control mice, IFNγ, IL-10 and IL-4 productions were reduced (P < 0·05) in response to PSC in mice fed and grafted with PSC. However, the IL-10/IFNγ and IL-4/IFNγ ratios increased (P < 0·04) in mice fed and subsequently grafted with PSC. These effects of feeding PSC on ‘secondary’ cytokine production were not detected when mice were fed PSC only after graft with PSC instead of prefeeding (Fig. 3c).

Effect of feeding PSC on humoral immunity

Compared to mice fed PBS, the production of IgM, IgG, IgG1, IgG2a and IgG2b directed against pig cells could not be detected in the sera of mice fed only with PSC (Fig. 4). Compared to mice fed PBS and grafted with PSC, levels of IgM against pig cells were increased (Fig. 4a; P < 0·04) in those fed PSC and subsequently grafted with PSC. IgG levels against pig cells were decreased (Fig. 4a; P < 0·04) in the same mice fed and grafted with PSC. More precisely, IgG2a and IgG2b levels were lower in mice fed PSC (P < 0·01), while IgG1 levels were not different in the two groups. These effects of prefeeding PSC were durable as they were observed throughout the experiment (up to 10 weeks after the last feeding and 8 weeks after grafting). The effects of feeding PSC on IgM, IgG or IgG isotypes were not detected when mice were fed PSC only after graft with PSC instead of prefeeding (Fig. 4b).

Fig. 4.

Fig. 4

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Humoral responses to pig cells in NOD mice fed with pig spleen cells (PSC). (a) Daily administrations of 25 × 106 PSC or PBS were given for 11 days. In some mice, antibody responses were tested after feeding PSC (○, n = 4) or PBS (□, n7= 4) alone. Other mice were analysed after feeding PSC (•, n = 5) or PBS (■, n = 6) followed by graft of PSC. Serum IgM, IgG, IgG1, IgG2a and IgG2b levels were tested by ELISA for up to 8 weeks after graft (10 weeks after the last feeding), using desiccated PSC (see Materials and methods). Each point represents the mean index of optical density (O.D.) ± s.e.m. * P < 0·04 and ** P < 0·01 indicate significant differences between mice fed with PSC or PBS. (b) Humoral responses to PSC when mice were fed PSC only after i.p. graft with PSC (instead of the prefeeding shown in (a)).

Generation of active transferable cells by feeding PSC

As compared to transfer with splenocytes from animals fed PBS, transfer with splenocytes from mice fed PSC increased the intensity of primary splenocyte proliferation in recipients in response to PSC (P < 0·02) or PIC (P < 0·04) (Fig. 5a). This enhancing effect of transfer on ‘primary’ proliferation was not decreased when recipients were irradiated prior to transfer (Fig. 5a).

Fig. 5.

Fig. 5

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Effect of adoptive transfer of splenocytes from mice fed pig spleen cells (PSC) on ‘primary’ and ‘secondary’ proliferations of recipient mice splenocytes in response to PSC or pig islet cells (PIC) in vitro. Daily oral administrations of 25 × 106 PSC were given for 11 days to NOD mice. Splenocytes were isolated 1 day after the last feeding and transferred three times (days 0, 3 and 14) i.v. to recipient mice (70 × 106 cells/transfer/mouse). In vitro‘primary’ proliferation (a) of 0·5 × 106 splenocytes from recipient mice (n = 14 per group) to 60 000 PSC or PIC was then tested on day 23 after the first transfer. In some experiments, recipient mice (n = 7) were irradiated prior to transfers. For analysis of ‘secondary’ proliferation (b), recipient mice (n = 10 in each group) were then i.p.-grafted (15 × 106 PSC on days 6 and 16 after the first transfer), and splenocyte proliferation of recipient mice was tested on day 23 after the first transfer. The results indicate the mean ± s.e.m. of the stimulation index (SI) of [3H]thymidine uptake, calculated as the ratio of the mean count with stimulator pig cells to the mean basal count. * P < 0·04 and ** P < 0·02 indicate significant differences between mice transferred with splenocytes from PSC-or PBS-fed mice.

Compared to mice grafted with PSC after transfer with splenocytes from donors fed PBS, mice grafted with PSC after transfer from PSC-fed donors displayed decreased (P < 0·04) ‘secondary’ proliferation to PSC or PIC (Fig. 5b). The increased ‘primary’ and decreased decreased ‘secondary’ splenocyte proliferations observed in mice receiving three injections of spleen cells from fed mice was also observed in recipient mice given a single transfer (data not shown). Compared to mice grafted with PSC after transfer with splenocytes from PBS-fed mice, IFNγ production was reduced whereas IL-10 production was increased in response to PSC in mice grafted with PSC after transfer with splenocytes from PSC-fed mice (Fig. 6). Thus, the IL-10/IFNγ ratio increased fourfold in mice grafted with PSC after being transferred with splenocytes from PSC-fed mice (Fig. 6).

Fig. 6.

Fig. 6

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Effect of transfer of splenocytes from mice fed pig spleen cells (PSC) on cytokine secretions from recipient mice in response to PSC. Daily oral administrations of 25 × 106 PSC were given for 11 days to NOD mice. Splenocytes were isolated 1 day after the last feeding and transferred three times (days 0, 3 and 14) i.v. to recipient mice (70 × 106 cells/transfer/mouse). IL-10 and IFNγ secretions were analysed 7 days after the second i.p. exposure in mice (n = 2 in each group) prefed PBS or PSC and i.p.-grafted (15 × 106 PSC on days 6 and 16 after the first transfer). The results indicate the mean ± s.e.m. production of IFNγ and IL-10 by 0·5 × 106 splenocytes alone (□) or co-incubated for 48 h with 60 000 pig cells (■). The mean ± s.e.m. of the IL-10/IFNγ ratio is also calculated.

DISCUSSION

We have reported previously that NOD mice fed with PSC displayed increased ‘primary’ proliferation of splenocytes in response to PSC or PIC [1] and that splenocytes from control mice inhibited basal and stimulated insulin releases from PIC in vitro, whereas this aggression was reduced during co-incubation with splenocytes from mice fed with PSC. By analogy with the down-regulation of systemic responses through oral administration of autoantigens [4–7], particularly in the NOD mouse [8–10], and of allogeneic cells [11–15], we suggested that feeding mice with PSC could have been conducive to active suppression. However, this preliminary study lacked several types of data.

In the present study, NOD mice fed with PSC and then exposed parenterally to PSC not only displayed increased ‘primary’ splenocyte proliferation but also subsequent lower ‘secondary’ splenocyte proliferation in response to PSC or PIC than control mice. Thus, oral administration of pig cells induced mechanisms characterized by a biphasic pattern with early activation and increased primary xenogeneic cellular reactions to pig cells, followed by decreased secondary responsiveness. Decreased secondary responsiveness may have been due to active suppression, as demonstrated for down-regulation of systemic immune responses by oral administration of autoantigens [4–7], particularly in the NOD mouse [8–10], and allogeneic cells [11–15]. However, it may also have resulted from the death or anergy of peripheral antigen-specific T cells, as demonstrated, for example, to account for the biphasic pattern of proliferative unresponsiveness to melin basic protein (MBP) after oral administration of MBP [21]. Several results suggest, albeit indirectly, that decreased ‘secondary’ proliferation in the present study was due to suppression.

First, our preliminary study had suggested that enhanced ‘primary’ splenocyte proliferation in response to PSC or PIC was related both to the number of daily feedings and of cells per feeding, showing an increase up to the highest number of cells per feeding tested. However, this highest number was only 25 × 106, so that an even higher number of fed PSC might induce hyporesponsiveness, i.e. anergy, described with high doses of orally administered antigens in other systems [22]. In the present study, the highest number of cells per feeding was increased to a very high level (200 × 106) without inducing hyporesponsiveness. Thus, feeding PSC in this model did not induce anergy, regardless of the number of cells per feeding.

Secondly, the treatment of mice with cyclophosphamide reduced both the enhancing effect on ‘primary’ splenocyte proliferation and the decrease in ‘secondary’ proliferation. Even though cyclophosphamide has complex effects, it is known to depress suppressor mechanisms in the NOD mouse [16]. Thus, the effects of this drug in our study suggest that feeding pig cells could have induced suppressor mechanisms which in turn could have reduced the subsequent response resulting from parenteral exposure to pig cells.

Thirdly, while IgM levels were increased, whole IgG levels against pig cells were decreased in mice fed and grafted with PSC. More precisely, IgG2a and IgG2b levels were lower, whereas IgG1 levels did not differ. Thus, oral administration of PSC also had a durable effect in modifying humoral reactions against pig cells. The observed modifications of whole IgG and isotypes suggest that feeding PSC reduced the Th1 stimulation induced by subsequent parenteral exposure. This is consistent with a shift of the Th2/Th1 balance towards a greater Th2 influence and the resulting shift of Ig isotypes following oral administration of antigens in other models [7,23]. Additionally, these results tend to confirm induction of suppression through the feeding of PSC. Induction of anergy or deletion would rather have induced a decrease of all Ig classes and isotypes.

The effects of feeding PSC on less intense Th1 stimulation, greater Th2 influence and induction of ‘secondary’ suppression were also confirmed by cytokine production during splenocyte proliferation in response to pig cells. During ‘primary’ proliferation, the IL-10/IFNγ and IL-4/IFNγ ratios were increased in mice fed with PSC, even though a slight increase of IFNγ production was also detected, confirming that not only Th2 but also Th1 cells were induced, as reported already in models of antigen feeding [24]. During ‘secondary’ proliferation, IFNγ production was more decreased than that of IL-10 and IL-4, so that the IL-10/IFNγ and IL-4/IFNγ ratios were increased. This confirms that feeding PSC induced greater Th2 influence during subsequent parenteral exposure to pig cells.

Additionally, feeding PSC induced active cellular mechanisms which were adoptively transferable. Transfer involving splenocytes from PSC-fed mice increased the intensity of ‘primary’ proliferation of splenocytes from recipient mice in response to PSC or PIC. This enhancing effect was not decreased when recipient mice were irradiated prior to transfer. Overall, transferred cells reproduced the effects of feeding PSC in recipient mice subsequently exposed in vivo to PSC, i.e. a reduction of ‘secondary’ splenocyte proliferation. The transfer of splenocytes from mice fed PSC also reproduced the effects of feeding PSC on cytokine profile during subsequent parenteral exposure to pig cells, in that the IL-10/IFNγ ratio in response to PSC tended to increase. Depletion and enrichment of cell subsets before transfer experiments could help indicate which cell populations were responsible for the effect.

Thus, the present study tends to suggest that induction of active suppression is a component of the mechanisms induced in mice fed PSC. In experimental systems distinct from xenoreaction, antigen feeding has given rise to CD8+ suppressor T (Ts) cells in gut-associated lymphoid tissue (GALT) and spleen [25,26], with the support of class II restricted Th cells [27]. APC from the intestinal lamina propria activate CD8+ T cells preferentially [28], and the lamina propria contains CD8+ T cells of naive phenotype [29], which indicates that GALT could be committed to the generation of Ts cells. In addition, CD4+ T cells of GALT can be activated following antigen feeding, which might be conducive to activation of CD8+ Ts cells [30]. In fact, antigen feeding modifies the Th1/Th2 ratio of CD4+ Th cells. Th2 cells secreting IL-4 and IL-10 [31, as well as a regulatory subset secreting transforming growth factor-β (TGF-β) [32], are mainly induced, even though the production of Th1 cytokines, e.g. IFN-γ, has also been demonstrated [24]. Our preliminary results [1] suggested that the mechanisms induced by feeding xenogeneic cells are consistent with these conclusions. In mice fed PSC, the blocking of MHC class II restricted CD4+ T cells led to abolition of ‘primary’ proliferation, while the blocking of CD8+ led to a less significant effect. However, a significant increase in the blocking effect of CD8 cells was noted in mice fed PSC. This might indicate that CD4 cells could have been conducive to activating CD8 Ts cells.

Additionally, the results of the present study also confirm that feedin]g pig cells induced cell activation which could then lead to suppression. A similar two-stage process occurred following oral administration of autoantigens [33,34]. This two-stage process induced by feeding, together with ‘secondary’ decreased splenocyte proliferation and an increased IL-1002F;IFNγ ratio, as well as decreased IgG production and isotype composition, could be detected in our study only when feeding of PSC preceded parenteral exposure. This suggests that different mechanisms are induced in GALT, depending on whether the xenoantigen/GALT interaction is or is not the first contact of mice with these antigens. This is also consistent with studies reporting that tolerance following oral administration of autoantigens was not easy to induce when autoantigen feeding followed parenteral injection of pathogenetic antigens [35,36].

The effects of feeding PSC or of transferring splenocytes from PSC-fed mice on increased ‘primary’ and decreased ‘secondary’ proliferation were not only observed with respect to PSC but also in response to PIC. Thus, these effects were specific for the species providing the fed cells, which could be of interest from a clinical point of view. It was not determined which pig antigen(s) of the fed spleen cells was (were) responsible for the observed effect. By analogy with the down-regulation of response to major histocompatibility complex (MHC) antigens by orally administered allogeneic cells [11–15], it may be speculated that molecules of the pig MHC shared by the fed cells were involved since they represent major targets for xenogeneic cellular recognition [37]. This hypothesis could be tested by feeding pig MHC antigens instead of whole cells to mice. Finally, it is also of interest that the effects of feeding xenogeneic cells were durable since they were detectable for at least 3 months after feeding or transfers.

The present study confirms the notion that oral administration of pig cells constitutes a novel approach to modifying xenogeneic cellular response in the mouse. It induces active transferable mechanisms, characterized by a biphasic pattern with early increased ‘primary’ xenogeneic cellular reactions to both PSC and PIC, followed by decreased ‘secondary’ responsiveness. Decreased responsiveness, concomitant with a shift of the Th1/Th2 balance towards greater Th2 influence, may have been due to active suppression, even though induction of anergy or deletion cannot be excluded. In fact, the two mechanisms are not exclusive; for example, apoptosis in antigen-activated T cells could be triggered by suppressor cells [38]. More generally, this study supports the concept that feeding of cells from xenogeneic species is not without effects on the immune system. Depending on the combination of parameters, feeding xenoantigens could probably induce opposite effects, i.e. stimulation or suppression of immune responses. It is difficult to extrapolate our results to man and to suppose that oral administration of xenoantigens could have application for xenografts. It is also uncertain whether the presence of pork products in the human diet could have an immunological impact.

Acknowledgments

We are grateful to Mrs M. Ouary and Mr C. Chevallier for their excellent collaboration. This work was supported by grants from the Pays de la Loire and Brittany regions. Pig islets were isolated from SPF pigs bred by a collaboration with AFSSA (Ploufragan, France, Dr Ph. Vannier).

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