Approaches to optimize immunosuppression after liver transplantation

__Abstract__ Since its advent 51 years ago, liver transplantation (LT) has progressed from an experimental treatment to an accepted therapeutic modality that has reversed the gloomy prognosis of end stage liver disease. The great success of LT is for the major part due to calcineurin inhibitors (CNI) like Cyclopsorin A and Tacrolimus. These are powerful immunosuppressants (IS), which prevent graft rejection and are the cornerstone of posttransplant patient management. The introduction of Cyclosporine in March 1980 marked great clinical advance for graft and patient survival. Following the introduction of cyclosporine in LT, one-year patient survival more than doubled (from approximately 33 to 68%), with an increment to above 80% nowadays. (www.eltr.org). Although immunosuppressive regimens are indispensible therapeutics for LT recipients, the downside is their major adverse effects on the long-term. The obligatory lifelong use of these drugs increases patients’ morbidity by increasing their susceptibility for infections, cancer, cardiovascular diseases, kidney failure and de-novo diabetes, which impair long-term survival and quality of life after transplantation. These adverse effects are largely caused by the non-specificity of the immunosuppressive medications. In addition CNI also exert undesirable side effects on vasoactive compounds resulting in vasculopathy, and on LDL receptor bile acid synthesis from cholesterol and lipoprotein lipase activity, resulting in dyslipidemia

Detailed Kinetics of the Direct Allo-Response in Human Liver Transplant Recipients: New Insights from an Optimized Assay

Conventional assays for quantification of allo-reactive T-cell precursor frequencies (PF) are relatively insensitive. We present a robust assay for quantification of PF of T-cells with direct donor-specificity, and establish the kinetics of circulating donor-specific T cells after liver transplantation (LTx). B cells from donor splenocytes were differentiated into professional antigen-presenting cells by CD40-engagement (CD40-B cells). CFSE-labelled PBMC from LTx-recipients obtained before and at several time points after LTx, were stimulated with donor-derived or 3rd party CD40-B cells. PF of donor-specific T cells were calculated from CFSE-dilution patterns, and intracellular IFN-γ was determined after re-stimulation with CD40-B cells. Compared to splenocytes, stimulations with CD40-B cells resulted in 3 to 5-fold higher responding T-cell PF. Memory and naïve T-cell subsets responded equally to allogeneic CD40-B cell stimulation. Donor-specific CD4+ and CD8+ T-cell PF ranged from 0.5 to 19% (median: 5.2%). One week after LTx, PF of circulating donor-specific CD4+ and CD8+ T cells increased significantly, while only a minor increase in numbers of T cells reacting to 3rd party allo-antigens was observed. One year after LTx numbers of CD4+ and CD8+ T cells reacting to donor antigens, as well as those reacting to 3rd party allo-antigens, were slightly lower compared to pre-transplant values. Moreover, CD4+ and CD8+ T cells responding to donor-derived, as well as those reacting to 3rd party CD40-B cells, produced less IFN-γ. In conclusion, our alternative approach enables detection of allo-reactive human T cells at high frequencies, and after application we conclude that donor-specific T-cell PF increase immediately after LTx. However, no evidence for a specific loss of circulating T-cells recognizing donor allo-antigens via the direct pathway up to 1 year after LTx was obtained, underscoring the relative insensitiveness of previous assays

Comparison of IFN-γ production by T cells in CFSE-MLR before and 1 year after LTx.

<p>CFSE-labeled PBMC from 5 LTx-patients were stimulated with donor-derived or 3^rd party CD40-B cells, and re-stimulated with the same allo-antigens at day 5 of culture for 24 hours. During the last 15 hours Brefeldin A was added, and CFSE-dilution and intracellular IFN-γ was determined at day 6. Depicted are the percentages of T cells producing IFN-γ in response to donor-derived CD40-B cells and 3^rd party-derived CD40-B cells, and the RR of IFN-γ producing T cells. No statistically significant differences were observed in comparing RR at 1 year after LTx versus before LTx (p≥0.44).</p

Expansion and differentiation of B cells using L-CD40L cells.

<p>A. Human splenocytes or PBMC were co-cultured with L-CD40L cells, IL-4 and CsA. Within 3 weeks, a 10⁴ fold increase of the numbers of B cells was obtained with a purity of >95%. T cells did not expand due to the presence of CsA. The current graph represents expansion from PBMC, splenocytes gave similar results. B. Cell surface marker expression on CD40-B cells harvested at day 12 of culture. CD40L-expanded B cells were activated (CD38⁺) and expressed the co-stimulatory molecules CD80 and CD86, and MHC class-II, indicating that they became professional APC. However, they did not become plasma cells, since they lacked CD138. Solid lines represent the expression of surface markers at day 0; dotted lines represent the expression at day 12 after culture. C. Donor splenocytes contained about 50% of HLA-DR⁺ APC, however these lacked CD80 and showed low expression of CD86.</p

Relative contributions of naïve and memory T cells to the allo-response in CFSE-MLR upon stimulation with CD40-B cells.

<p>CFSE-labelled naïve CD3⁺CD45RA⁺and CD3⁺CD45RO⁺ memory T cells were isolated from PBMC of healthy individuals by flowcytometric sorting, and stimulated with allogeneic CD40-B cells for 6 days. A. Purity of CD3⁺CD45RO⁺ and CD3⁺CD45RA⁺ T cells after sorting. B. PF of CD3⁺CD45RO⁺ and CD3⁺CD45RA⁺ T cells proliferating upon stimulation with allogeneic CD40-B cells. Depicted are means ± SD of data from two independent experiments, each with 3 replicates.</p

Analysis of CFSE-dilution patterns by ModFit® software.

<p>ModFit-derived CFSE-pattern of CD3⁺ T cells after 6 days of stimulation with allogeneic CD40-B cells. The software draws peaks within the CFSE-histogram centered on halving intensity values from the parental peak CFSE-intensity. Non-divided T cells are CFSE^high and are depicted blue, while the peaks left from it correspond to divided cells. Based on the enumerated proportions of T cells detected in each generation ( = % under each peak) and the total number of CD3⁺ T cells analyzed by the flowcytometer, the program calculates the absolute numbers of daughter T cells in each generation. The numbers of precursors which gave rise to the daughter cells are extrapolated by dividing the absolute numbers of T cells in each generation by 2ⁿ, in which n stands for the division cycle ( = absolute # of precursors). The PF is calculated by dividing the numbers of precursors from generation 2 onwards ( = B) by the total number of precursors in all generations including the parent peak and generation 1 ( = A). In this example the calculated PF =  (362,302 – (337,620 + 8925)/362,302) ×100 = 4.3%. The fit of the derived Gaussian peaks in the CFSE-dilution pattern is indicated by the reduced chi-square value. Values below 5 were considered as a good fit according to the manufacturer's instructions, and were included in the analysis.</p

Kinetics of allogeneic T-cell responses to CD40-B cell stimulation.

<p>CFSE-labeled PBMC from a healthy individual were cultured for 7 days with allogeneic splenocyte-derived CD40-B cells. A. CFSE-dilution patterns of CD3⁺, CD4⁺, CD8⁺ T cells were analyzed daily, and PF were calculated with ModFit® software. B. Intracellular granzyme B and perforin expression were analyzed daily in CD8⁺ T cells. Depicted are the percentages of CD8⁺CFSE^low T cells that expressed granzyme B and/or perforin. C. Absolute numbers CD3⁺, CD4⁺ and CD8⁺ T cells during co-culture of CFSE-labelled PBMC with allogeneic CD40-B cells. Vital PBMC were counted daily using trypan blue, and proportions of CD3⁺, CD4⁺ and CD8⁺ T cells were analyzed by flowcytometry. Absolute numbers of T-cell subsets were calculated by multiplying the number of vital PBMC with the proportions of T cells obtained from flowcytometric analysis.</p

Longitudinal course of circulating donor-specific T-cell precursor frequencies in LTx-recipients.

<p>CFSE-labeled PBMC of 13 LTx-recipients obtained before transplantation (Pre), or at 1 week (W1), 1 month (M1), 3 months (M3) or 1 year (Y1) after transplantation, were co-cultured for 6 days with donor-derived splenic CD40-B cells, 3^rd party splenic CD40-B cells or autologous PBMC-derived CD40-B cells. PF of CD3⁺, CD4⁺ and CD8⁺ T cells responding to these stimulators were determined. Recipients and donors differed on the average in 1.5 HLA-AB alleles and in 1.7 HLA-DR alleles. Third-party stimulators were mismatched with recipients in 1.7 HLA-AB alleles and 1.8 HLA-DR alleles. Donor-derived and 3^rd party-derived stimulator cells differed on the average at 1.8 HLA-AB loci and 1.5 HLA-DR loci. A. Numbers of T-cell precursors responding to donor-derived CD40-B cells increased significantly 1 week after transplantation in all T-cell subsets, followed by a decrease to values below pre-transplant levels. B. Non-specific variations in allo-responses were determined by stimulating patient PBMC with third-party spleen-derived CD40-B cells. Changes in PF of CD4⁺ and CD8⁺ T cells responding to 3^rd party CD40-B cells between subsequent time points were generally smaller compared to those in donor-specific PF. C. Donor-specific responses were calculated by dividing PF responding to donor-alloantigen by third-party PF to obtain the relative responses (RR). RR increased significantly 1 week after transplantation in all T-cell subsets, followed by a significant decrease at 1 year after LTX. D. Comparison of relative CD3⁺, CD4⁺ and CD8⁺ T-cell responses (RR) of 18 LTx-recipients before and 1 year after LTx. Five additional patients were analyzed, and their data were added to the data of the 13 patients shown in C. Donor-derived and 3^rd party-derived stimulator cells used in assaying the 5 additional patients were fully MHC mismatched.</p