Myeloid dendritic cells (MDC) are amongst the most potent antigen-presenting cells, and as such play a critical role in the initiation and direction of immune responses.1, 2 Under normal circumstances, most peripheral tissues contain immature MDC whose function is uptake and processing of antigens. Upon antigenic stimulation, they migrate toward the paracortex of the draining lymph nodes (LN) and spleen. During this migration MDC mature and acquire a strong T-cell stimulatory capacity.1–3 Animal experiments indicate that after transplantation donor dendritic cells (DC) migrate from the graft into the recipient's regional LN and spleen.4–7 In a murine liver transplantation model donor DC were even traced in the recipient's spleen 7 months after transplantation.8
It is generally accepted that donor MDC are the main instigators of acute rejection after solid organ and tissue transplantation due to their potent capacity to stimulate recipient T-cell responses against the graft.4, 5, 7 Unlike experimental mouse heart or skin allografts, mouse liver allografts are accepted across major histocompatibility complex barriers and induce donor-specific tolerance without antirejection therapy.8 Interstitial DC of rodent liver are comparatively rare cells that are located predominantly in portal areas and, very occasionally, in sinusoids.9, 10 Freshly isolated murine liver DC are predominantly immature, expressing surface major histocompatibility complex but few co-stimulatory molecules.11–13 In addition, DC generated in vitro from murine liver leukocytes exhibited an immature phenotype.13–15 Injecting these in vitro propagated liver-derived DC pretransplantation has been shown to induce donor-specific hyporeponsiveness.16, 17 However, a marked increase of DC numbers and maturation state in liver allografts by donor pretreatment with the hematopoietic growth factor Flt3 ligand results in acute rejection.17, 18 This implies a crucial function for liver DC in regulating the balance between rejection and tolerance to liver grafts.
In spite of the ample knowledge on the properties of rodent liver DC, less is known about DC in human livers. Scarceness of human liver tissue, the relative low frequency of DC within human liver tissue, and, until recently, the paucity of DC-specific reagents that recognize all maturation stages of human DC have hampered their investigation. DC-like cells have been detected immunohistochemically in the portal triads of the human liver.19 Recently it has been shown that MDC migrating from human liver tissue in vitro have a relative immature phenotype. These MDC have a high expression of major histocompatibility complex classes II and CD86, but a relatively low expression of CD80 and CD83.20 Nevertheless, hepatic MDC have the capacity to mature in vivo, since MDC from hepatic LN have a mature phenotype, with higher expressions of these molecules than MDC from skin/muscle draining LN.21, 22
Until now no data on freshly isolated human resident liver DC are available, since it is difficult to obtain sufficient healthy human liver tissue for characterization of scarce cells like hepatic DC. It has been shown that substantial numbers of liver leukocytes readily detach from the donor liver during vascular perfusion of liver grafts before transplantation.23 If such perfusates contain substantial numbers of DC, these could be used to study human donor liver DC.
The aim of this study was to characterize MDC in human liver grafts. Since the second type of DC, plasmacytoid DC (PDC), are less efficient in antigen presentation, and recent data on their role in transplantation suggest that they facilitate organ graft survival,24 we have focused on the classical MDC. MDC were visualized by immunohistochemistry on liver cryosections using the anti-BDCA-1 monoclonal antibody (mAb), which is expressed on both immature and mature MDC. Donor liver mononuclear cells (MNC) were used for a limited analysis of the maturational status of liver MDC by flow cytometry. In addition, we found that large numbers of viable MDC were present in perfusates of human liver grafts obtained during ex vivo vascular perfusion pretransplantation. Perfusate MDC were used for further immunophenotypic and functional characterization of human donor liver MDC.
Wedge biopsies were obtained prior to transplantation from 10 liver grafts and used for isolation of liver leukocytes (n = 10) or were snap frozen for immunohistochemistry (n = 7). Perfusates were collected from 20 liver grafts that immediately after arrival in the Erasmus MC Transplant Center during the back-table procedure were perfused through the portal vein with 1 to 2 L of University of Wisconsin solution to remove residual blood from the vasculature. Immediately before transplantation the donor liver was again perfused ex vivo through the portal vein with 200 up to 500 mL of human albumin solution under hydrostatic pressure, and the perfusate was collected from the vena cava. Hepatic LN (n = 12) were obtained from the hepatoduodenal ligament of donor livers during preparation of the liver pretransplantation. Blood samples were collected from 3 of the multiorgan donors and from 12 healthy volunteers. Nine hepatic LN were obtained from the same donors as perfusates; 5 of 10 liver wedge biopsies were obtained from the same donors as perfusates. The multiorgan donor blood samples were obtained from the same donors as both perfusates and hepatic LN. The Ethics Committee of the Erasmus MC approved the study protocol, and informed consent of each patient was obtained.
The following mAbs were used: IgG1-FITC, IgG1-PE, IgG1-APC, IgG1-PerCP-cy5.5, IgG2a-FITC, CD4-PerCP, CD19-FITC, CD20-PerCP, CD14-FITC, HLA-DR-FITC, CD86-APC, and streptavidin-APC from Becton Dickinson, Heidelberg, Germany; CD45-FITC, CD3-PE, CD56-APC, CD80-FITC, and anti-DC-LAMP-PE from Beckman Coulter Immunotech, Marseille, France; anti-BDCA-1 pure (= CD1c), anti-BDCA1-PE, anti-BDCA1-FITC, anti-BDCA2-FITC, CD19-microbeads, and anti-PE-microbeads from Miltenyi Biotec, Bergisch Gladbach, Germany; CD8-APC and CD45-RPE-Cy5 from DAKO, Glostrup, Denmark; CD83-APC from Caltag, Burlingame, CA; anti-CCR7-PE from R&D systems, Abingdon, UK.
To identify the location of MDC in donor livers pretransplantation, 5-μm cryostat sections from donor liver biopsies were stained with anti-BDCA-1 mAb. Optimal dilutions of mAbs were established in preliminary experiments by titration on human tonsil cryosections. Briefly, cryosections were fixed in acetone (10 minutes), after which endogenous peroxidase was blocked by incubation in citric acid/phosphate buffer solution (pH = 5.8) with 0.05% H2O2and 0.2% NaN3 (15 minutes, 20°C). The slides were washed with Tris-buffered saline (pH 7.4) twice, after which anti-BDCA-1 or IgG2a isotype-matched controls were applied in optimal concentrations in Tris-buffered saline (pH 7.4) supplemented with 0.01% Normal Human Plasma for 18 hours at 4°C. Binding of anti-BDCA-1 mAb was detected by incubation with conjugated goat anti-mouse Envision-PO (Envision™, DAKO, Glostrup, Denmark) for 30 minutes at room temperature and visualized using 3-amino-9-ethylcarbazole. The sections were counterstained with Mayers hematoxylin (blue) (Merck, Haarlem, The Netherlands).
Additionally, anti-BDCA-1 mAb was also used in double staining with CD19 mAb. Sections were first incubated with FITC-conjugated CD19 for 1 hour at room temperature, and subsequently with alkaline phosphatase-conjugated rabbit anti-FITC immunoglobulins (DAKO). Than slides were incubated overnight at 4°C with anti-BDCA-1, and binding was detected with PO-Envision. Visualization of alkaline phosphatase was performed by incubation in Fast Blue salt/ naphtol AS-BI phosphate solution supplemented with levamisole, giving a blue precipitate. Revelation of peroxidase was performed with 3-amino-9-ethylcarbazole, giving a red precipitate.
Six portal fields and 6 microscopic fields of 400× magnification in the parenchyma were analyzed, immunohistochemically positive cells were counted, and means with standard deviation (SD) were calculated.
Isolation of MNC
Cells were harvested from the perfusion fluid by centrifugation and resuspended in 30 mL of RPMI+ (RPMI 1640 with L-glutamine, Cambrex Bio Science, Verviers, Belgium). For isolation of single cells, the fresh liver tissue wedge biopsies were cut into small pieces that were incubated in RPMI+ supplemented with collagenase type IV (0.5 mg/mL; Gibco, Breda, The Netherlands) and DNase type I (0.02 mg/mL; Roche Diagnostics, Manheim, Germany) for 40 minutes at 37°C. Subsequently the tissue pieces were passed over a nylon mesh filter (200-μm-pore diameter) to obtain a single cell suspension. To remove hepatocytes the suspension was centrifuged at 360 rpm for 2 minutes and supernatant was collected. LN were also cut into small pieces and passed over a nylon mesh filter to obtain a single cell suspension. MNC were obtained from perfusates, single cell suspensions of wedge biopsies, and LN, and blood by Ficoll-Paque (Amersham Biosciences, Roosendaal, The Netherlands) density centrifugation. Cell viability was determined using trypan blue staining.
Maturation of Perfusate MDC
Perfusate MNC were routinely cultured in RPMI+ supplemented with 10% fetal calf syndrome (Hyclone, Logan, UT), pencillin (100 U/mL), and streptomycin (100 μg/mL; Gibco BRL Life Technologies, Breda, The Netherlands) for 24 hours at 37°C in the presence of 100 ng/mL lipopolysaccharide (LPS) (Sigma, Zwijndrecht, The Netherlands) in 24-well plates using 1 × 106 MNC per well. After 24 hours the cells were harvested and expression of maturation antigens and co-stimulatory molecules on MDC was determined using flow cytometric analysis.
Isolation of MDC from Blood and Perfusate MNC
For isolation of MDC 80 μL of PBS supplemented with 2 mmol/L ethylenediamine tetraacetic acid and 5 mg/mL bovine serum albumin (BSA), 100 μL of CD19 microbeads, and 20 μL of anti-BDCA1-PE were added per 100 × 106 MNC, and the cells were incubated for 15 minutes at 4°C. Hereafter, B-cells were depleted by separation over a large depletion column using a MidiMACS separation device (Miltenyi Biotec). The nonadherent cells were incubated for 15 minutes at 4°C with 50 μL of anti-PE microbeads and separated over a mini-separation column using a MiniMACS device, after which the adherent cells were washed out and enriched further by separation over a second MS column. Purity of the isolated MDC was determined by flow cytometry, and viability by trypan blue exclusion. Purity and viability of perfusate MDC were respectively 92 ± 10% and 89 ± 8%, and of blood MDC respectively 86 ± 10% and 83 ± 13%.
Flow cytometric analysis was used to immunophenotype the MNC. MNC were resuspended in PBS with human immunoglobulin G (IgG) (1.25 μg/mL Octagam; Octapharma, Langenfeld, Germany) to prevent aspecific binding of antibodies to Fc-receptors on DC. Per labeling 1 × 106 MNC were incubated with antibodies. The following antibody combinations were used to determine the different subsets of the MNC: CD45-FITC, CD3-PE, CD4-PerCP, and CD8-APC; CD45-FITC, CD3-PE, and CD56-APC; CD45-FITC, anti-BDCA1-PE, and CD20-PerCP; or anti-BDCA2-FITC and CD45-RPE-Cy5. MDC were defined as BDCA-1+ and CD20− cells and PDC as BDCA2+ cells. To determine the phenotype of MDC, MNC were labeled with the following antibody combinations: anti-BDCA1-PE and CD20-PerCP in combination with anti-HLA-DR-FITC, CD83-APC, CD80-FITC, and CD86-APC; or anti-BDCA1-FITC and CD20-PerCP in combination with anti-CCR7-PE, or in combination with intracellular staining with anti-DC-LAMP-PE. For the intracellular staining MNC were first incubated with anti-BDCA1-FITC and CD20-PerCP, then permeabilized and fixed with IntraPrep Permeabilization Reagent according to the manufacturer's protocol (Beckman Coulter Immunotech, Marseille, France) before adding the anti-DC-LAMP-PE. Cell death of the DC was determined using 7-amino-actinomycin-diacetate staining (BD Biosciences Pharmingen, San Diego, CA). Appropriate isotype-matched control antibodies were used. Optimal dilutions of all antibodies were established in preliminary experiments. The effects of collagenase and DNase treatment on cell surface expression of the above markers was tested on blood MDC and no alterations were observed (data not shown). The data were analyzed on a FACScalibur using Cellquest Pro software.
Allogeneic T-Cell Stimulatory Capacity of Purified MDC
Purified MDC were co-cultured at different concentrations (10, 5, 2.5, and 1.25 × 103 cells/200 μL) in a flat-bottom Costar culture plate (Costar, Cambridge, MA) with 1.5 × 105 purified T-cells from the blood of a healthy volunteer. After 5 days, cell proliferation was assessed by measuring the incorporation of [3H]-thymidine (Radiochemical Center, Amersham, Little Chalfont, UK). 0.5 μCi was added per well and cultures were harvested 18 hours later. Phytohemagglutinin (5 μg/mL; Murex, Paris, France) was added to the T -ells as a positive control. T-cells were purified by incubation of peripheral blood MNC with CD14-PE, anti-BDCA1-PE, and CD19 microbeads, and subsequently with anti-PE microbeads for 15 minutes at 4°C. T-cells were enriched by negative selection over a large separation column using a MidiMACS separation device (Miltenyi Biotec) and contained 78% CD3+ T-cells and 16% CD56+ cells.
Purified MDC were cultured at a concentration of 4 × 104 cells/200 μL in a 96-well flat-bottom culture plate in RPMI+ supplemented with 10% fetal calf syndrome (Hyclone, Logan, UT), pencillin (100 U/mL), streptomycin (100 μg/mL; Gibco BRL Life Technologies, Breda, The Netherlands), and granulocyte-macrophage colony-stimulating factor (500 U/mL; Leucomax, Novartis Pharma, Arnhem, The Netherlands) in the presence or absence of 1 μg/mL LPS for 24 hours at 37°C. After 24 hours, supernatants were harvested and the level of IL-10 was determined by specific sandwich enzyme-linked immunosorbent assay, using a mAb and recombinant cytokine standard from Biosource International (Camarillo, CA).
The Kruskal-Wallis and Mann-Whitney tests from SPSS version 11.0 were used to test whether differences between groups were statistically significant. The Wilcoxon signed ranks test from SPSS version 11.0 was used to test whether differences between groups of paired samples were statistically significant. A P of <0.05 was considered significant. All data are presented as means ± SD.
MDC in Human Donor Livers
MDC were visualized in wedge biopsies of donor livers (n = 7) by immunohistochemistry with anti-BDCA-1 mAb. BDCA-1+ cells were predominantly observed in the portal fields (average, 6.1 ± 2.3 cells per portal field), and only a few resided in the parenchyma (average, 0.6 ± 0.5 cells per microscopic field; P = 0.001) (Fig. 1A and B). Since BDCA-1 is also expressed on a subpopulation of B-cells,25 additional double stains with anti-BDCA-1 and CD19 mAb were performed (Fig. 1C and D). Portal fields contained on average 4.9 ± 6.0 BDCA-1+CD19− MDC per portal field, which is comparable to the number of BDCA-1+ cells observed in the single stains with anti-BDCA-1 mAb (P = 0.351). Both in portal tracts and parenchyma only few BDCA-1/CD19 double-positive B-cells were observed.
To characterize the MDC present in donor livers, MNC were isolated from donor liver wedge biopsies (n = 10). Liver MNC contained on average 1.0% (range, 0.2-2.2%) BDCA-1+CD20− MDC. The expression of CD83 and CD80 was determined on these DC (Fig. 2A). Donor liver MDC had an immature phenotype characterized by a relatively low expression of the maturation marker CD83, as well as the co-stimulatory molecule CD80 (Fig. 2B and C).
Composition of Perfusate MNC in Comparison with Donor Liver and Blood
The perfusates collected during the pretransplantation ex vivo albumin perfusions of the donor livers had an average volume of 378 ± 93 mL and contained 59 ± 36 × 106 MNC. The MNC present in the perfusates (99 ± 2%) were vital as determined by trypan blue staining. The CD4/CD8 ratio in perfusates (0.7 ± 0.4) was identical to that in the donor liver MNC (0.6 ± 0.1; P = 1.000), and both perfusate (P = 0.003) and donor liver (P = 0.014) MNC differed significantly from blood (3.6 ± 1.9) (Fig. 3). Also, the natural killer cell proportion, defined as the percentage of CD45+ cells expressing CD56, in perfusate was similar to that in liver MNC (respectively 44 ± 12% and 28 ± 16%; P = 0.413), but perfusate differed significantly from blood (13 ± 5%; P = 0.000). The observed similarity in CD4/CD8 ratio and percentage of natural killer cells between liver and perfusate and the significantly distinct ratios of these cells in blood strongly suggest that leukocytes present in perfusates are predominantly liver tissue derived.
DC in Perfusate, Hepatic LN, and Blood
Perfusate MNC contained on average 1.5% (range, 0.3-6.6%) MDC (= BDCA-1+CD20− cells), with a vitality of 97 ± 2% determined by 7-AAD staining. The total number of MDC that detached from donor livers during vascular perfusion pretransplantation was 0.9 × 106 (range, 0.11-4.5 × 106). Perfusate MNC contained a significantly higher percentage of MDC in comparison with LN (0.6%; range, 0.2-1.1%; P = 0.03), and MDC percentages in perfusates tended to be higher than in blood (0.7%; range, 0.2-1.3%; P = 0.11). In addition, perfusate MNC contained on average 0.9% (range, 0.2-2.6%) PDC (= BDCA-2+ cells), similar to LN (0.3%; range, 0.1-0.5%) and blood (0.5%; range 0.3-0.7%; P > 0.10). The total number of PDC that detached from donor livers during vascular perfusion pretransplantation was 0.5 × 106 (range, 0.1-1.3 × 106). The variation in volumes and numbers of MNC did not correlate with differences in cold ischemic times (Bosma, B.M., data not shown). Altogether, these data indicate that a high number of DC detach during perfusion from donor livers pretransplantation.
Maturation Markers, Co-stimulatory Molecules, and CCR7 on Perfusate MDC in Comparison with Liver, Blood, and Hepatic LN MDC
To determine whether MDC in liver perfusate are representative for resident liver MDC, the expression of CD80 and CD83 on perfusate and liver MDC from the same multiorgan donor were pairwise compared. The mean fluorescence intensities (MFI) (Fig. 4) were similar. Additionally, a comparison of the expression of CD80 was made between liver, perfusate, and blood MDC obtained from the same multiorgan donors. Similar numbers of liver and perfusate MDC expressed CD80 (6.7 ± 4.6% and 8.1 ± 6.4% respectively; P = 1.0), but a significantly lower number of blood MDC expressed CD80 (2.4 ± 1.1%; P < 0.03). These data indicate that perfusate MDC are liver MDC and that these MDC can be used to study the properties of hepatic MDC. To further investigate the maturation state of liver MDC, we compared the expression of the maturation markers CD14, DC-LAMP, CD83, and HLA-DR on perfusate MDC with blood and hepatic LN MDC. On average, 48% of the perfusate MDC and blood MDC expressed CD14, while on LN MDC the monocyte marker was almost absent (Fig. 5A). Expression of DC-LAMP was low on perfusate MDC, but intermediate between blood and LN MDC (Fig. 5B). Perfusate MDC had a significant lower expression of HLA-DR in comparison with LN MDC (Fig. 5C). There was no significant difference in CD83 expression between MDC derived from the different materials (Fig. 5D).
Analysis of the expression of the co-stimulatory molecules and the LN homing receptor CCR7 on MDC revealed that the expression of CD80 was low on perfusate MDC, and intermediate between blood and LN MDC (Fig. 5E). CD86 expression on blood and perfusate MDC was similar, but the expression on LN MDC was significantly higher (Fig. 5F). The observed differences between MDC from perfusates and hepatic LN were confirmed in paired comparisons of MDC from the same multiorgan donor (n = 9): CD80 (P = 0.012), CD86 (P = 0.004), CD14 (P = 0.016), and DC-LAMP (P = 0.063). MDC in blood and perfusate expressed significantly lower levels of CCR7 than LN MDC (P ≤ 0.014) (Fig. 6).
Maturation of Perfusate MDC In Vitro
To assess whether perfusate MDC are capable of maturing in vitro, perfusate MNC were incubated with 100 ng/mL LPS for 24 hours (n = 3). Twenty-four hours of incubation with LPS resulted in an increase of the expression of both maturation markers and co-stimulatory molecules on perfusate MDC (Fig. 7). Thus, MDC from liver grafts respond to a common bacterial danger signal.
IL-10 Production and Stimulation of Allogeneic T-Cells by Perfusate MDC Compared with Blood MDC
To determine whether perfusate MDC were able to stimulate allogeneic T-cell proliferation, graded numbers of perfusate MDC were cultured with allogeneic T-cells. The T-cell stimulatory capacity of freshly isolated perfusate MDC was compared to immature blood MDC. Perfusate MDC stimulated allogeneic T-cells, whereas blood MDC were not able to induce T-cell proliferation (Fig. 8).
Recently Goddard et al.20 showed that human liver MDC produce high levels of IL-10 in comparison with skin DC. To ascertain that perfusate MDC are liver derived, IL-10 production of perfusate MDC was compared with blood MDC. Both types of MDC did not produce IL-10 when cultured without a stimulus; however, when MDC were stimulated with 1 μg/mL, LPS perfusate MDC (n = 3) produced significantly more IL-10 than blood MDC (n = 7) (1800 ± 1875 pg/mL and 137 ± 45 pg/mL, respectively; P = 0.017).
To our knowledge this is the first study in which freshly isolated human liver MDC have been characterized by flow cytometric analysis. The obtained data indicate that MDC from human donor livers have a relatively low expression of the maturation marker CD83 and co-stimulatory molecule CD80, indicating that they are immature DC. These findings are consistent with data from murine studies showing that liver DC have a high expression of surface major histocompatibility complex class II, but few co-stimulatory molecules.11, 13 The immunochemical stains with anti-BDCA-1 mAb indicate that the majority of MDC reside in the portal fields of human donor livers.
The presence of liver-specific immune cells, namely, T-cells, natural killer cells, monocytes, and B-cells, in perfusates of donor livers has been previously reported.23 Detection of DC was probably not attempted in the previous study due to the lack of suitable DC-specific markers at that time. The present study demonstrates that perfusates contained on average 0.9 × 106 MDC and 0.5 × 106 PDC. Since PDC are less efficient in antigen presentation and first data on their role in transplantation suggest that they do not induce rejection,24 we have focused on the classical MDC.
Our data show that MDC present in perfusates were liver MDC and not blood MDC: First, perfusate MDC had a similar CD80 and CD83 expression as resident liver MDC. Furthermore, CD80 expression was slightly higher on perfusate MDC than on blood MDC. Second, perfusate MDC produced significantly higher amounts of IL-10 than blood MDC upon stimulation with LPS. High IL-10 production is also characteristic for liver MDC, as shown by Goddard et al.20 Third, perfusate MDC were able to stimulate allogeneic T-cell proliferation in contrast to freshly isolated blood MDC. Fourth, the MNC in perfusates had a low CD4/CD8 ratio and a high percentage of natural killer cells, which is indicative for liver-derived cells.26 Containing high numbers of MDC, perfusates are a rich source for the study of donor liver MDC.
Using perfusate, donor liver MDC were further immunophenotyped. The maturation state of hepatic MDC was compared with immature blood MDC and mature hepatic LN MDC.1, 21, 27 Perfusate MDC, like blood MDC, have a high expression of CD14, which indicates that these MDC are derived either from CD14+ immature MDC in the blood, from CD14+ bone marrow precursors, or from monocytes.28 The expression of DC-LAMP, a lysosomal protein that is up regulated in mature MDC,29 and HLA-DR is significantly lower on perfusate MDC than LN MDC. In addition, the co-stimulatory molecules CD80 and CD86 are expressed significantly lower on perfusate MDC than LN MDC. CCR7, normally expressed by mature MDC that have acquired the ability to home to secondary lymphoid tissue,30 is also expressed significantly lower on perfusate MDC than LN MDC. Together these results demonstrate that donor liver MDC have an immature phenotype.
Furthermore, the majority of isolated perfusate MDC are vital and functional since they have the capacity to produce high amounts of IL-10, to stimulate allogeneic T-cell proliferation, and to mature in vitro when exposed to the maturation stimulus LPS. These data show that perfusate MDC are functionally intact and can be used as a source for functional studies on donor liver MDC.
Another important implication of our study is that resident liver DC leave the liver graft via the vasculature during ex vivo perfusion. As the lymph vessels are transected during transplantation, we suppose that DC also leave the transplanted graft after reperfusion via the vasculature and migrate into the blood circulation. However, this does not exclude another possible migration route, namely, via the transected lymph vessels into the peritoneum of the recipient, and then via the lymphatics and blood circulation to the secondary lymphoid organs.
About one million MDC, which have the potential to become immunogenic, detach from the donor liver pretransplantation via vascular perfusion. We hypothesize that posttransplantation similar numbers of MDC, and probably even higher, will detach from the liver allograft, migrate into the recipient, and subsequently induce acute rejection. Similar numbers of MDC are sufficient for induction of immunomodulatory processes; from a clinical trial for treatment of advanced metastatic melanoma (stage VI), it is known that intravenous injection of as little as 0.9 × 106 MDC can induce a tumor-specific immune response.31 Thus, the numbers of donor-derived MDC that detach during perfusion from donor livers are expected to be high enough to induce an allogeneic immune response in the recipient. Ischemia/reperfusion injury could elicit the maturation of donor liver DC in vivo. Upon arrival in the recipient's LN or spleen these donor MDC are thought to present allogeneic major histocompatibility complex molecules to the recipient T-cells, which are recognized via the direct pathway, causing acute rejection.32 From animal studies it is known that mature donor liver DC are indeed capable of inducing acute rejection and that, on the contrary, immature DC can induce tolerance.14, 16, 17, 33
Since MDC in donor livers as well as in perfusates are immature, they may still be sensitive to pharmacological targeting, thereby preventing acute rejection.34 Pharmacological pretreatment of the donor liver during the cold storage could be a promising new method to manipulate MDC to induce tolerance, and isolation of this cell fraction from perfusates would allow in vitro testing of such a model.
In conclusion, human donor livers contain exclusively immature MDC that detach in high numbers from the liver graft during perfusion before transplantation. These vital MDC have the capacity to stimulate allogeneic T-cells and to mature upon activation, and could therefore play a major role in the induction of acute rejection.
We are grateful to Dr. L. van der Laan for his scientific input and to Dr. T.C.K. Tran and Dr. J.N.M. IJzermans for collecting the liver wedge biopsies, perfusates, and LN.