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reperfusion injury inhibits differentiation of dendritic cells derived from bone marrow monocytes in rats
Life Sciences 78 (2006) 1121 – 1128 www.elsevier.com/locate/lifescie
Renal ischemia/reperfusion injury inhibits differentiation of dendritic cells derived from bone marrow monocytes in rats Chih-Jen Wu a,b, Joen-Rong Sheu b, Han-Hsiang Chen a,c,d, Hui-Fen Liao e,f, Yuh-Cheng Yang e, Stone Yang g, Yu-Jen Chen b,e,h,i,* a Division of Nephrology, Mackay Memorial Hospital, Taipei, Taiwan Graduate Institute of Medical Science, Taipei Medical University, Taipei, Taiwan c Mackay Medicine, Nursing and Management College, Taipei, Taiwan d Taipei Nursing College, Taipei, Taiwan e Department of Medical Research, Mackay Memorial Hospital, Taipei, Taiwan Department of Molecular Biology and Biochemistry, National Chiayi University, Chiayi, Taipei, Taiwan g Division of Urology, Mackay Memorial Hospital, Taipei, Taiwan h Radiation Oncology, Memorial Hospital, Taipei, Taiwan i Graduate Institute of Sports Coaching Science, Chinese Culture University, Taipei, Taiwan b
f
Received 5 February 2005; accepted 21 June 2005
Abstract Dendritic cells (DCs) are impacted by surgical injury, exercise, and other physiological stressors. This study aims to determine whether renal I/ R injury affects 1) the differentiation of myeloid DCs from bone marrow monocytes (BMMos) and the maturation and activation state of these DCs and 2) DC infiltration of kidney. Sprague – Dawley rats were subjected to I/R injury or sham-operated. Creatinine clearance was monitored daily during the 14 d of reperfusion that followed the ischemic insult. At 2 and 14 d of reperfusion, the following were assessed 1) properties of BMMo-derived DCs (i.e., the amount of generated DCs, differentiation state markers [CD11c, CD80, CD86, and Ia], and functional state [MLR and amount of IL-12 produced]), and 2) the presence of DCs in the kidney. Numbers of BMMo-derived DCs were significantly decreased in the I/ R injured group (compared with the sham-operated group) at 2 d but not 14 d. A comparison of the their functionality found mixed lymphocyte response [MLR] and IL-12 production were similar in the two groups at both time points. Also, immunohistochemistry showed infiltrating DCs in the outer medulla of the I/R injured kidney at 2 d but not 14 d of reperfusion. Thus, I/R stress reduces the number of DCs differentiated from BMMos but not the functional activity of these DCs. This decrease may reflect a stress-induced downshift in the capacity of BMMos to differentiate into DCs and a parallel upshift in the capacity of DCs to infiltrate the kidney. D 2005 Elsevier Inc. All rights reserved. Keywords: Myeloid dendritic cells; Bone marrow-derived dendritic cells; Ischemia – reperfusion injury; Differentiation
Introduction The most potent antigen-presenting cells (APCs) are dendritic cells (DCs). These are migratory cells derived mainly from monocytes and therefore referred to as ‘‘myeloid DCs’’. DCs reside in most tissues and organs where they exert a mobile sentinel function for incoming antigens. Upon captur* Corresponding author. Department of Radiation Oncology, Mackay Memorial Hospital, 92 Chung San North Road, Section 2, Taipei 104, Taiwan. Tel.: +886 2 28094661; fax: +886 2 28096180. E-mail address: [email protected] (Y.-J. Chen). 0024-3205/$ - see front matter D 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.lfs.2005.06.043
ing antigen, myeloid DCs mature and migrate to secondary lymphoid organs where the presented antigen stimulates T cell activation. In vitro, myeloid DCs stimulate T-helper 1 (Th1) or Th2 responses depending on the maturation conditions (Rissoan et al., 1999; Vieira et al., 2000). These conditions may profoundly influence immunological outcome and the role that DCs play in this outcome. Ischemia/reperfusion (I/R) injury has been associated with ischemic acute renal failure (Bonventre and Weinberg, 2003), which is a major cause of native kidney and allograft dysfunction (Lieberthal and Nigam, 1998). DCs appear to play a central role in the initiation of an adaptive immune response
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to I/R-induced tissue injury. In the case of renal I/R injury, reactive oxygen species contribute to both the development of injury (De Vecchi et al., 1998) and maturation of DCs. Hydrogen peroxide has been shown 1) to upregulate MHC class II and co-stimulatory molecules CD40 and CD86 on peripheral blood monocyte-derived DCs (such cells are capable of stimulating T cell proliferation [Rutault et al., 1999]) and 2) to stimulate the production of IL-8 and TNFa by peripheral blood monocyte-derived DCs in a dose-dependent manner, suggesting that DCs could contribute to innate immunity in response to oxidative stress (Verhasselt et al., 1998). Moreover, injury of transplantation itself was found to result in the infiltration of recipient MHC class II-positive leukocytes (some of which were DCs) into the transplanted kidney, suggesting that the transplant recipient rats were mounting an adaptive immune response (Penfield et al., 1999). The monocyte-derived DC response to renal I/R injury has not been defined or clearly distinguished from the DC response to injury due to the renal transplant surgery itself. Also unclear is whether the DC response has a role in the injury phase or in the regeneration process after I/R. The objective of this paper was to determine the effect of I/R injury on DC differentiation from BMMos, the maturation and activation state of these DCs, and DC infiltration of kidney. Methods and materials Rats Adult Sprague – Dawley rats (200 – 250 g) obtained from the National Animal Center in Taiwan were cared for, and all experiments conducted, according to the guidelines of the National Science Council of the Republic of China (NSC, 1997). Induction of renal ischemia/reperfusion (I/R) Animals were randomly allocated into four groups (two I/R groups [1 and 3] and two sham groups [2 and 4]): Group 1 (N = 12) experienced ischemia and 2 d of reperfusion; Group 2 (N = 12), no ischemia and 2 d of reperfusion; Group 3 (N = 12), ischemia and 14 d of reperfusion; and Group 4 (N = 12), no ischemia and 14 d of reperfusion. Micraneurysm clamps were used to close each renal artery and vein for 40 min. Renal function measurement Tail vein blood and 24-h urine samples were collected from all rats housed in metabolic cages. Plasma and urine creatinine was determined using a standard colorimetric method (Bonsnes and Taussky, 1945). Generation of monocyte-derived dendritic cells from bone marrow Generation of large numbers of immature and mature dendritic cells was according to the procedure of Talmor et
al. (1998). Bone marrow (BM) collected from rat femurs and humeri were cultured in complete medium (RPMI 1640 with 10% fetal bovine serum [FBS], 1% glutamine, 50 AM 2mercaptoethanol, and 20 Ag/ml gentamycin) for 2 h. After the nonadherent cells were aspirated, the adherent cells were washed with Hank’s Balanced Salt Solution (HBSS) and incubated in complete medium supplemented with recombinant granulocyte – macrophage colony-stimulating factor (rGMCSF; 5 ng/ml; R&D Systems, Minneapolis, MN, USA) and IL-4 (5 ng/ml; R&D Systems). The medium was replaced on d 3, and the DCs were stimulated on d 6 with lipopolysaccharide (LPS) (5 Ag/ml). On d 7, the loosely adherent cells were harvested and the supernatants were collected and stored at 20 -C for IL-12 determination. Expression of surface markers was quantified by flow cytometry. Generation of monocyte-derived dendritic cells from peripheral blood Rat mononuclear cells were obtained from whole blood collected via cardiac puncture into heparin (10 U/ml). Buffy coats were collected and underlayered with Lymphoprep (product no. 1053980, Nycomed Pharma AS, Oslo, Norway), and centrifuged at 1430 g for 20 min. The peripheral blood mononuclear cells (PBMCs) were obtained from the interface between the plasma and Ficoll solution, treated with ammonium chloride to lyse the remaining erythrocytes, washed twice with phosphate buffered saline (PBS) to remove the platelets, resuspended in culture medium, and transferred into six-well plates (1.2 107 cells in 3 ml/well). After incubation (37.8 -C, 2 h), nonadherent cells were removed by gentle washing with PBS, suspended in medium (final concentration 2 106 cells/ml), and incubated in 100-mm tissue culture dishes (Becton-Dickinson, Franklin Lakes, NJ, USA) for 2 h. The nonadherent cells were gently washed out with warm medium, and to generate DCs from the remaining adherent cells, R10 medium (which consisted of RPMI1640 supplemented with 10% FBS, 5 ng/ml of rat recombinant granulocyte – macrophage colony-stimulating factor [rGMCSF], and rat IL-4 [rIL-4]) was added. The culture conditions were 5% CO2, 37 -C, and media change every 3 d. Both suspended and adherent cells were propagated. Clusters of adherent cells with dendritic morphology were detached using PBS + 3 mM EDTA. The remaining strongly adherent cells were discarded. After 6 d of culture, the immature DCs were collected and stimulated with 5 Ag/ml of lipopolysaccharide (LPS) for 24 h to generate mature DCs. The purity of the DC population was checked by flow cytometry. Dendritic cell morphology DCs (80 Al of 1 106) were loaded and spun in a Cytospin centrifuge at 1000 rpm for 5 min. The slides were dried at least 1 h. The cells were stained with Liu A staining solution for 45 s and then 2 Liu B staining solution for 45 s. The slides were washed in tap water, air dried, and examined under a microscope.
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Flow cytometry
Fig. 1. Light microscopic view of Wright – Giemsa-stained veiled cells generated from peripheral blood adherent cells in the presence of rGM-CSF and rIL-4 and then LPS. Magnification 1000.
Fluorescence activated cell sorting (FACS) analysis was performed as described elsewhere (Sallusto and Lanzavecchia, 1994; Verhasselt et al., 1998). Briefly, BMDC preparations (1 105) were incubated for 30 min with mouse monoclonal antibodies specific for rat CD11c, CD80, CD86, and HLA-DR (Serotec, Oxford, England), washed twice with phosphate buffered saline (PBS)– 10% FBS, incubated with fluorescein isothiocyanate-(FITC)-conjugated goat anti-rat IgG (30 min on ice; ICN, Costa Mesa, CA, USA), washed twice with PBS – 10% FBS, incubated with propidium iodide (100 nM, 30 min on ice; Sigma, St. Louis, MO, USA), washed twice with PBS – 10% FBS, and analyzed by means of the FACSCalibur flow cytometry system (Becton-Dickinson, Mountain View, CA, USA) and CELLQuest software.
Fig. 2. Flow cytometric analysis of myeloid DC markers (CD11c, CD80, CD86, and Ia) demonstrating decreases in myeloid DCs at A) 2 d but not at B) 14 d of reperfusion. FACs profiles in C) show no difference in per-cell differentiation marker fluorescence between the two groups at both 2 and 14 d of perfusion. Significant differences in marker levels at *p < 0.05 on d 2 only.
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Fig. 2 (continued).
Mixed lymphocyte reaction assay T cells were removed from peripheral blood or spleens using plastic adherence. About 1 105 DCs were co-cultured with 1 106 T cells in culture dishes for 7 d. BrdU (10 AM; 5bromo-2-deoxyuridine) was added to the culture medium on d 6. After 18 h, the cells were collected and resuspended in 200 Al of PBS with 10% FBS. One ml of 70% ethanol ( 20 -C) was slowly added to the cells, and then the mixture was incubated (on ice, 30 min). After washing in PBS – 10% FBS, 1 ml of 2N HCl-Triton X100 was added slowly to the cells and incubated (30 min, room temperature). The cells were centrifuged and resuspended in 1 ml of 0.1M Na2B4O7 to neutralize the acid. After washing in PBS – 10% FBS, the cells were resuspended in 200 Al of 0.5% Tween 20 –1% BSA – PBS and incubated with anti-BrdU FITC
(Becton-Dickinson, San Jose, CA, USA) for 30 min at room temperature. The cells were washed with PBS – 10% FBS, and propidium iodide was added to determine the percentage of dead cells. After washing in PBS –10% FBS, the cells were analyzed using a flow cytometer. Table 1a Generation of myeloid DCs from BMMos
I/R 2 d Sham 2 d I/R 14 d Sham 14 d
BMMos ( 105/rat)
DCs (105/rat)
Recovery rate (%)
3647.6 T 341.5 3827.2 T 253.3 4678.0 T 221.5 5156.7 T 172.4
55.3 T 16.2* 90.8 T 22.7 68.3 T 1.2 79.5 T 1.9
1.4 T 0.4* 2.5 T 0.8 1.5 T 0.3 1.6 T 0.4
DCs: dendritic cells; BMMos: bone marrow monocytes. Recovery rate = (DCs BMMos) 100%. * p < 0.05 as compared with the sham 2 d group.
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Measurement of IL-12 IL-12 release from myeloid DCs and plasma IL-12 were measured using an ELISA kit (Biosource Inc., Camarillo, CA, USA) according to the manufacturer’s protocol. Immunohistochemistry of dendritic cells in kidney All rats were perfused via the left cardiac ventricle with about 300 ml of 0.002% heparin sodium in 0.9% sodium
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chloride solution. The kidneys were removed and fixed by immersion in 4% paraformaldehyde (PFA)/PBS at 4 -C overnight, transferred to 30% sucrose/PBS at 4 -C overnight, embedded in Tissue-Tek\ O.C.T compound (Miles Inc., Elkhart, IN, USA), and stored at 80 -C until ready for cryostat sectioning. Sections (5 Am thick) mounted on slides were incubated with mouse monoclonal antibodies specific to CD11c+, CD80, or CD86 (Serotec, Oxford, England) and then with biotin-conjugated goat anti-mouse immunoglobulin and horseradish peroxidase (HRP)-conjugated streptavidin
Fig. 3. DCs stained by antibodies to CD11c A) and D), CD80 B) and E), and CD86 C) and F) are shown in the outer medulla of the kidney at 2 d (A – C) but not at 14 d (D – F) of reperfusion.
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Table 1b Generation of PBDCs from PBMos
I/R 2 d Sham 2 d
Table 2b DC IL-12 secretion by BMDCs
PBMos (105/rat)
PBDCs (105/rat)
Recovery rate (%)
619.0 T 61.5 665.8 T 52.9
21.0 T 2.9 13.5 T 2.9
3.9 T 0.3* 2.1 T 0.5
DCs: dendritic cells; PBMo: peripheral blood monocytes. Recovery rate = (DCs PBMo) 100%. * p < 0.05 as compared with the sham 2 d group.
IL-12 (pg/ml) I/R 2 d Sham 2 d I/R 14 d Sham 14 d
105.1 T 32.1* 217.4 T 51.3 174.2 T 99.6 153.6 T 81.6
IL-12 (pg/105 DCs) 34.6T6.5 38.2 T 12.7 29.1 T17.3 37.6 T 19.4
* p < 0.05 as compared with the sham 2d group.
(DAKO, Glostrup, Denmark). Aminoethylcarbazole was used as chromogenic substrate, and the presence of a red precipitate was identified as positive staining.
differentiation markers on BMMo-derived DCs (Fig. 2C) was not significantly different at both 2 and 14 d. Dendritic cell infiltration of the kidney
Statistical analysis Data are expressed as the mean T SE. All comparisons were made by analysis of variance followed by Dunnett’s t-test using the Statview software package for the Macintosh computer (Abacus Concepts, Berkeley, CA, USA). Results
Immunohistochemical analysis of CD11c, CD80, and CD86 labeling of DCs in kidney sections from I/R injured rats showed DCs are present in the outer medulla at 2 d (Fig. 3A – C) but not 14 d of reperfusion (Fig. 3D – F). DCs were not seen in kidney sections of sham-operated rats at either time. Generation of peripheral blood monocyte-derived DCs at 2 d of reperfusion
Changes in renal function after I/R injury Renal dysfunction, indicated by elevated plasma creatinine levels and decreased calculated creatinine clearance, was observed on d 0 through 4 (peak day 1—0.32 T 0.12 ml/min) of reperfusion in rats subjected to I/R (Gr 1 and Gr 3) compared to sham-operated rats (Gr 2 and Gr 4) (data not shown). Generation of bone marrow monocyte-derived DCs Myeloid DCs generated from BM cells cultured in the presence of rGM-CSF and rIL-4 for six days and in the presence of LPS for one additional day were recovered as a population of nonadherent or loosely adherent clustered and veiled cells (Fig. 1) and recognized by flow cytometry as a population of CD11c-CD80-CD86-Ia-bearing cells (Fig. 2A and B). In preliminary experiments, no significant difference in DC differentiation was observed between d 1 and 5 of reperfusion (data not shown). However, the number of mature BMMo-derived DCs was significantly lower at 2 d after reperfusion in I/R rats than in sham-operated rats (Gr 1 vs. Gr 2), whereas this number was not significantly different at 14 d (Gr 3 vs. Gr 4) (Table 1a) and the per-cell expression of Table 2a Mixed lymphocyte response of peripheral blood (PB) and splenic Tlymphocytes to stimulator bone marrow dendritic cells (BMDCs) BMDCs from rats treated with:
Control
PB T-cells
Control
Splenic T-cells
I/R 2 d Sham 2 d I/R 14 d Sham 14 d
9.6 T 1.6 9.6 T 1.6 4.5 T 1.0 4.5 T 1.0
32.5 T 4.2 31.5 T 4.6 26.0 T 6.9 21.9 T 6.8
3.1 T 0.4 3.1 T 0.4 4.0 T 1.0 4.0 T 1.0
20.3 T 4.4 22.4 T 4.1 25.3 T 9.4 20.7 T 6.2
The results are expressed as percent of BrdU (5-bromo-2-deoxyuridine)positive cells and means T SEM for N = 12. No significant differences between I/R- and sham-treated animals are shown.
Because BMMo differentiation to DCs decreased at 2 d at the same time that DCs increased in the outer medulla of the kidney, we thought that PBMos might also be responders at 2 d of reperfusion to ischemic injury. Hence, PBMo-derived DCs were counted and found to be significantly higher at 2 d (Table 1b). Mixed lymphocyte reaction stimulated by BMMo-derived DCs was unaffected by I/R injury Whether the source of T cells was peripheral blood or spleen (Table 2a), no significant differences in MLR proliferation were found between I/R rats and sham-operated rats. IL-12 production of BMMo-derived DCs was unaffected by I/R injury Though myeloid DCs from I/R rats produced considerably less IL-12 by 2 d of reperfusion than myeloid DCs from shamoperated rats, the difference reflected merely the decrease in cell numbers. There was no significant difference in the per 105 cell release of IL12 between the groups (Gr 1 vs. Gr 2 and Gr 3 vs. Gr 4; Table 2b). In plasma, IL-12 levels were undetectable in all groups (detection level, 0 –500 pg/ml). Discussion Our study suggests that I/R injury 1) significantly decreases the number of myeloid DCs differentiated from BMMos at 2 d but not at 14 d of reperfusion 2) has no effect on functional activity (i.e., no change in DC-stimulated MLR and in IL-12 synthesis of individual DCs), and 3) increases the number of DCs in the kidney at 2 d. These changes may reflect inhibition of DC differentiation from BMMos and either stimulation of
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DC differentiation from Mos in the kidney or migration of DCs to the kidney during the acute phase of I/R injury. I/R injury was also found to increase the number of DCs differentiated from peripheral blood monocytes (PBMos) at 2 d. The difference between the differentiability of monocytes in BM and monocytes in PB suggests that these cells represent two distinct subpopulations. In support of this possibility, Sunderkotter et al. have demonstrated in mice that immature Ly6Chigh monocytes (phenotypically BMMos) immigrate from the BM to the circulation, and that during inflammation they predominate for > 2 d in the circulation where they gradually mature to Ly6Clow monocytes (phenotypically PBMos). Under steady state conditions, the predominant monocytes in the circulation were the more mature Ly6Clow cells, whereas under inflammatory conditions, the most abundant were immature Ly6Chigh (the phenotype in bone marrow) and less mature Ly6Cmedium cells (Sunderkotter et al., 2004). These findings imply that the primary distinction between monocyte subsets is their maturation state and that environmental factors such as inflammatory conditions determine subset predominance. Similarly, another study found that maturation state and therefore DC subset is influenced by environmental factors: the presence of CD40 ligand (which accelerates maturation of DCs) and phagocytosis of bacteria and yeasts (which favors development of less mature DCs) (Rosenzwajg et al., 2002). Thus, microenvironmental influences appeared to direct monocyte maturation and subset development. Our study also showed I/R injury resulted in DC infiltration of the outer medulla of the kidney after 2 d but not 14 d of reperfusion. We can only speculate that the transience of DC appearance reflects either DC entry followed by DC exodus from the kidney or further maturation of mature DCs, e.g., into endothelial-like cells (Havemann et al., 2003). Analogously, Kennedy and Abkowitz found that the more mature monocyte subpopulation of their monocyte transplant landed in various organs 3 d after transplantation and that by 7 d, these cells had differentiated into macrophages in situ. At 3 d, cells were found as clusters and individual cells. By 7 and 14 d, the clusters had increased in size and the cells expressed the macrophage antigen F4/80, suggesting that further replication and differentiation had occurred (Kennedy and Abkowitz, 1998). Our results are consistent with this idea that bone marrow or peripheral blood DCs or DC progenitors migrate to kidney via peripheral blood following I/R injury. Also, Sunderkotter et al. observed in mice experimentally depleted of monocytes the presence of repopulating macrophages and DCs in peripheral organs (particularly spleen and liver) 4 d after monocyte depletion (Sunderkotter et al., 2004). The functional role of DCs that settle in the renal medulla as a result of I/R injury is unclear. This will require further investigation. As for the fate of these DCs, our study found that the DC response to I/R injury (decreased differentiation from BMMos, increased differentiation from PBMos, and infiltration of the renal outer medulla, which is the area susceptible to I/R injury [Brezis et al., 1984]) occurred only during the acute phase (d 2) of renal I/R injury. This finding suggests 1) that DC differentiation from BMMos may shift to PBMos, then DCs
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may migrate from PB to the kidney and 2) that this shift and migration are stress-induced. Because our study focused on differentiation and did not track the migration of DCs, we do not know whether the decreased DCs in BM is due to migration to PB and then to kidney. Penfield et al. have shown that injury to the kidney during transplantation is sufficient to recruit recipient MHC class II leukocytes (some of which are DCs) into the transplant (Penfield et al., 1999). Similarly, Loi et al. showed, in a mouse model, that liver I/R injury itself induces DC maturation, migration, and preferential production of inhibitory cytokines (Loi et al., 2004). However, they failed to show whether the DCs in the liver parenchyma were recruited in situ or were derived from circulating precursors. Because PBDCs, BMDCs, and DCs infiltrating the renal medulla may be different subpopulations and because these cells may interact with and stimulate different T cell populations, comparison of these cells is uncertain and the problem requires further investigation. Two aspects of the MLR technique used in this study should be mentioned because they may be regarded as limitations. First, the technique was modified (BrdU [a nonradioactive analogue of thymidine] rather than 3H-thymidine was measured (Gratzner, 1982)), and second, a mixed T cell population rather than bead-purified T cells were added. In summary, the upshift in PBMo differentiation to DCs, downshift in BMMo differentiation to DCs, and infiltration of the kidney at 2 d of reperfusion may be part of an important pathophysiologic acute stress response to renal I/R injury. Acknowledgments We thank Gabriel Cheng, Ming-Ling Hsu, Hui-Ju Shieh, Hung-Jen Liao and Ching-Pin Lin for their technical help and writing assistance. This work was supported by research grants from the National Science Council (NSC 93-2413-H-195-001) and Mackay Memorial Hospital Medical Research Fund (MMHE93006 and MMH9372). References Bonventre, J.V., Weinberg, J.M., 2003. Recent advances in the pathophysiology of ischemic acute renal failure. Journal of the American Society of Nephrology 4, 2199 – 2210. Bonsnes, R.M., Taussky, H.M., 1945. The colorimetric determination of creatinine by Jaffe’s reaction. Journal of Biological Chemistry 158, 581 – 591. Brezis, M., Rosen, S., Silva, P., Epstein, F.H., 1984. Renal ischemia: a new perspective. Kidney International 26 (4), 375 – 383. De Vecchi, E., Lubatti, L., Beretta, C., Ferrero, S., Rinaldi, P., Galli Kienle, M., Trazzi, R., Paroni, R., 1998. Protection from renal ischemia – reperfusion injury by the 2-methylaminochroman U83836E. Kidney International 54 (3), 857 – 863. Gratzner, H.G., 1982. Monoclonal antibody to 5-bromo- and 5-iododeoxyuridine: a new reagent for detection of DNA replication. Science 218, 474 – 475. Havemann, K., Pujol, B.F., Adamkiewicz, J., 2003. In vitro transformation of monocytes and dendritic cells into endothelial like cells. Advances in Experimental Medicine and Biology 522, 47 – 57. Kennedy, D.W., Abkowitz, J.L., 1998. Mature monocytic cells enter tissues and engraft. Proceedings of the National Academy of Sciences of the United States of America 95 (25), 14944 – 14949.
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Lieberthal, W., Nigam, S.K., 1998. Acute renal failure. I. Relative importance of proximal vs. distal tubular injury. American Journal of Physiology 275, F623 – F631. Loi, P., Paulart, F., Pajak, B., Nagy, N., Salmon, I., Moser, M., Goldman, M., Flamand, V., 2004. The fate of dendritic cells in a mouse model of liver ischemia/reperfusion injury. Transplantation Proceedings 36 (5), 1275 – 1279. Penfield, J.G., Wang, Y., Li, S., Kielar, M.A., Sicher, S.C., Jeyarajah, D.R., Lu, C.Y., 1999. Transplant surgery injury recruits recipient MHC class II-positive leukocytes into the kidney. Kidney International 56 (5), 1759 – 1769. Rissoan, M.C., Soumelis, V., Kadowaki, N., Grouard, G., Briere, F., de Waal Malefyt, R., Liu, Y.J., 1999. Reciprocal control of T helper cell and dendritic cell differentiation. Science 283 (5405), 1183 – 1186. Rosenzwajg, M., Jourquin, F., Tailleux, L., Gluckman, J.C., 2002. CD40 ligation and phagocytosis differently affect the differentiation of monocytes into dendritic cells. Journal of Leukocyte Biology 72 (6), 1180 – 1189. Rutault, K., Alderman, C., Chain, B.M., Katz, D.R., 1999. Reactive oxygen species activate human peripheral blood dendritic cells. Free Radical Biology & Medicine 26 (1 – 2), 232 – 238.
Sallusto, F., Lanzavecchia, A., 1994. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. Journal of Experimental Medicine 179, 1109 – 1118. Sunderkotter, C., Nikolic, T., Dillon, M.J., Van Rooijen, N., Stehling, M., Drevets, D.A., Leenen, P.J., 2004. Subpopulations of mouse blood monocytes differ in maturation stage and inflammatory response. Journal of Immunology 172 (7), 4410 – 4417. Talmor, M., Mirza, A., Turley, S., Mellman, I., Hoffman, L.A., Steinman, R.M., 1998. Generation or large numbers of immature and mature dendritic cells from rat bone marrow cultures. European Journal of Immunology 28 (3), 811 – 817. Verhasselt, V., Goldman, M., Willems, F., 1998. Oxidative stress up-regulates IL-8 and TNF-alpha synthesis by human dendritic cells. European Journal of Immunology 28, 3886 – 3890. Vieira, P.L., de Jong, E.C., Wierenga, E.A., Kapsenberg, M.L., Kalinski, P., 2000. Development of Th1-inducing capacity in myeloid dendritic cells requires environmental instruction. Journal of Immunology 164 (9), 4507 – 4512.
Renal ischemia/reperfusion injury inhibits differentiation of dendritic cells derived from bone marrow monocytes in rats Chih-Jen Wu a,b, Joen-Rong Sheu b, Han-Hsiang Chen a,c,d, Hui-Fen Liao e,f, Yuh-Cheng Yang e, Stone Yang g, Yu-Jen Chen b,e,h,i,* a Division of Nephrology, Mackay Memorial Hospital, Taipei, Taiwan Graduate Institute of Medical Science, Taipei Medical University, Taipei, Taiwan c Mackay Medicine, Nursing and Management College, Taipei, Taiwan d Taipei Nursing College, Taipei, Taiwan e Department of Medical Research, Mackay Memorial Hospital, Taipei, Taiwan Department of Molecular Biology and Biochemistry, National Chiayi University, Chiayi, Taipei, Taiwan g Division of Urology, Mackay Memorial Hospital, Taipei, Taiwan h Radiation Oncology, Memorial Hospital, Taipei, Taiwan i Graduate Institute of Sports Coaching Science, Chinese Culture University, Taipei, Taiwan b
f
Received 5 February 2005; accepted 21 June 2005
Abstract Dendritic cells (DCs) are impacted by surgical injury, exercise, and other physiological stressors. This study aims to determine whether renal I/ R injury affects 1) the differentiation of myeloid DCs from bone marrow monocytes (BMMos) and the maturation and activation state of these DCs and 2) DC infiltration of kidney. Sprague – Dawley rats were subjected to I/R injury or sham-operated. Creatinine clearance was monitored daily during the 14 d of reperfusion that followed the ischemic insult. At 2 and 14 d of reperfusion, the following were assessed 1) properties of BMMo-derived DCs (i.e., the amount of generated DCs, differentiation state markers [CD11c, CD80, CD86, and Ia], and functional state [MLR and amount of IL-12 produced]), and 2) the presence of DCs in the kidney. Numbers of BMMo-derived DCs were significantly decreased in the I/ R injured group (compared with the sham-operated group) at 2 d but not 14 d. A comparison of the their functionality found mixed lymphocyte response [MLR] and IL-12 production were similar in the two groups at both time points. Also, immunohistochemistry showed infiltrating DCs in the outer medulla of the I/R injured kidney at 2 d but not 14 d of reperfusion. Thus, I/R stress reduces the number of DCs differentiated from BMMos but not the functional activity of these DCs. This decrease may reflect a stress-induced downshift in the capacity of BMMos to differentiate into DCs and a parallel upshift in the capacity of DCs to infiltrate the kidney. D 2005 Elsevier Inc. All rights reserved. Keywords: Myeloid dendritic cells; Bone marrow-derived dendritic cells; Ischemia – reperfusion injury; Differentiation
Introduction The most potent antigen-presenting cells (APCs) are dendritic cells (DCs). These are migratory cells derived mainly from monocytes and therefore referred to as ‘‘myeloid DCs’’. DCs reside in most tissues and organs where they exert a mobile sentinel function for incoming antigens. Upon captur* Corresponding author. Department of Radiation Oncology, Mackay Memorial Hospital, 92 Chung San North Road, Section 2, Taipei 104, Taiwan. Tel.: +886 2 28094661; fax: +886 2 28096180. E-mail address: [email protected] (Y.-J. Chen). 0024-3205/$ - see front matter D 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.lfs.2005.06.043
ing antigen, myeloid DCs mature and migrate to secondary lymphoid organs where the presented antigen stimulates T cell activation. In vitro, myeloid DCs stimulate T-helper 1 (Th1) or Th2 responses depending on the maturation conditions (Rissoan et al., 1999; Vieira et al., 2000). These conditions may profoundly influence immunological outcome and the role that DCs play in this outcome. Ischemia/reperfusion (I/R) injury has been associated with ischemic acute renal failure (Bonventre and Weinberg, 2003), which is a major cause of native kidney and allograft dysfunction (Lieberthal and Nigam, 1998). DCs appear to play a central role in the initiation of an adaptive immune response
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to I/R-induced tissue injury. In the case of renal I/R injury, reactive oxygen species contribute to both the development of injury (De Vecchi et al., 1998) and maturation of DCs. Hydrogen peroxide has been shown 1) to upregulate MHC class II and co-stimulatory molecules CD40 and CD86 on peripheral blood monocyte-derived DCs (such cells are capable of stimulating T cell proliferation [Rutault et al., 1999]) and 2) to stimulate the production of IL-8 and TNFa by peripheral blood monocyte-derived DCs in a dose-dependent manner, suggesting that DCs could contribute to innate immunity in response to oxidative stress (Verhasselt et al., 1998). Moreover, injury of transplantation itself was found to result in the infiltration of recipient MHC class II-positive leukocytes (some of which were DCs) into the transplanted kidney, suggesting that the transplant recipient rats were mounting an adaptive immune response (Penfield et al., 1999). The monocyte-derived DC response to renal I/R injury has not been defined or clearly distinguished from the DC response to injury due to the renal transplant surgery itself. Also unclear is whether the DC response has a role in the injury phase or in the regeneration process after I/R. The objective of this paper was to determine the effect of I/R injury on DC differentiation from BMMos, the maturation and activation state of these DCs, and DC infiltration of kidney. Methods and materials Rats Adult Sprague – Dawley rats (200 – 250 g) obtained from the National Animal Center in Taiwan were cared for, and all experiments conducted, according to the guidelines of the National Science Council of the Republic of China (NSC, 1997). Induction of renal ischemia/reperfusion (I/R) Animals were randomly allocated into four groups (two I/R groups [1 and 3] and two sham groups [2 and 4]): Group 1 (N = 12) experienced ischemia and 2 d of reperfusion; Group 2 (N = 12), no ischemia and 2 d of reperfusion; Group 3 (N = 12), ischemia and 14 d of reperfusion; and Group 4 (N = 12), no ischemia and 14 d of reperfusion. Micraneurysm clamps were used to close each renal artery and vein for 40 min. Renal function measurement Tail vein blood and 24-h urine samples were collected from all rats housed in metabolic cages. Plasma and urine creatinine was determined using a standard colorimetric method (Bonsnes and Taussky, 1945). Generation of monocyte-derived dendritic cells from bone marrow Generation of large numbers of immature and mature dendritic cells was according to the procedure of Talmor et
al. (1998). Bone marrow (BM) collected from rat femurs and humeri were cultured in complete medium (RPMI 1640 with 10% fetal bovine serum [FBS], 1% glutamine, 50 AM 2mercaptoethanol, and 20 Ag/ml gentamycin) for 2 h. After the nonadherent cells were aspirated, the adherent cells were washed with Hank’s Balanced Salt Solution (HBSS) and incubated in complete medium supplemented with recombinant granulocyte – macrophage colony-stimulating factor (rGMCSF; 5 ng/ml; R&D Systems, Minneapolis, MN, USA) and IL-4 (5 ng/ml; R&D Systems). The medium was replaced on d 3, and the DCs were stimulated on d 6 with lipopolysaccharide (LPS) (5 Ag/ml). On d 7, the loosely adherent cells were harvested and the supernatants were collected and stored at 20 -C for IL-12 determination. Expression of surface markers was quantified by flow cytometry. Generation of monocyte-derived dendritic cells from peripheral blood Rat mononuclear cells were obtained from whole blood collected via cardiac puncture into heparin (10 U/ml). Buffy coats were collected and underlayered with Lymphoprep (product no. 1053980, Nycomed Pharma AS, Oslo, Norway), and centrifuged at 1430 g for 20 min. The peripheral blood mononuclear cells (PBMCs) were obtained from the interface between the plasma and Ficoll solution, treated with ammonium chloride to lyse the remaining erythrocytes, washed twice with phosphate buffered saline (PBS) to remove the platelets, resuspended in culture medium, and transferred into six-well plates (1.2 107 cells in 3 ml/well). After incubation (37.8 -C, 2 h), nonadherent cells were removed by gentle washing with PBS, suspended in medium (final concentration 2 106 cells/ml), and incubated in 100-mm tissue culture dishes (Becton-Dickinson, Franklin Lakes, NJ, USA) for 2 h. The nonadherent cells were gently washed out with warm medium, and to generate DCs from the remaining adherent cells, R10 medium (which consisted of RPMI1640 supplemented with 10% FBS, 5 ng/ml of rat recombinant granulocyte – macrophage colony-stimulating factor [rGMCSF], and rat IL-4 [rIL-4]) was added. The culture conditions were 5% CO2, 37 -C, and media change every 3 d. Both suspended and adherent cells were propagated. Clusters of adherent cells with dendritic morphology were detached using PBS + 3 mM EDTA. The remaining strongly adherent cells were discarded. After 6 d of culture, the immature DCs were collected and stimulated with 5 Ag/ml of lipopolysaccharide (LPS) for 24 h to generate mature DCs. The purity of the DC population was checked by flow cytometry. Dendritic cell morphology DCs (80 Al of 1 106) were loaded and spun in a Cytospin centrifuge at 1000 rpm for 5 min. The slides were dried at least 1 h. The cells were stained with Liu A staining solution for 45 s and then 2 Liu B staining solution for 45 s. The slides were washed in tap water, air dried, and examined under a microscope.
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Flow cytometry
Fig. 1. Light microscopic view of Wright – Giemsa-stained veiled cells generated from peripheral blood adherent cells in the presence of rGM-CSF and rIL-4 and then LPS. Magnification 1000.
Fluorescence activated cell sorting (FACS) analysis was performed as described elsewhere (Sallusto and Lanzavecchia, 1994; Verhasselt et al., 1998). Briefly, BMDC preparations (1 105) were incubated for 30 min with mouse monoclonal antibodies specific for rat CD11c, CD80, CD86, and HLA-DR (Serotec, Oxford, England), washed twice with phosphate buffered saline (PBS)– 10% FBS, incubated with fluorescein isothiocyanate-(FITC)-conjugated goat anti-rat IgG (30 min on ice; ICN, Costa Mesa, CA, USA), washed twice with PBS – 10% FBS, incubated with propidium iodide (100 nM, 30 min on ice; Sigma, St. Louis, MO, USA), washed twice with PBS – 10% FBS, and analyzed by means of the FACSCalibur flow cytometry system (Becton-Dickinson, Mountain View, CA, USA) and CELLQuest software.
Fig. 2. Flow cytometric analysis of myeloid DC markers (CD11c, CD80, CD86, and Ia) demonstrating decreases in myeloid DCs at A) 2 d but not at B) 14 d of reperfusion. FACs profiles in C) show no difference in per-cell differentiation marker fluorescence between the two groups at both 2 and 14 d of perfusion. Significant differences in marker levels at *p < 0.05 on d 2 only.
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Fig. 2 (continued).
Mixed lymphocyte reaction assay T cells were removed from peripheral blood or spleens using plastic adherence. About 1 105 DCs were co-cultured with 1 106 T cells in culture dishes for 7 d. BrdU (10 AM; 5bromo-2-deoxyuridine) was added to the culture medium on d 6. After 18 h, the cells were collected and resuspended in 200 Al of PBS with 10% FBS. One ml of 70% ethanol ( 20 -C) was slowly added to the cells, and then the mixture was incubated (on ice, 30 min). After washing in PBS – 10% FBS, 1 ml of 2N HCl-Triton X100 was added slowly to the cells and incubated (30 min, room temperature). The cells were centrifuged and resuspended in 1 ml of 0.1M Na2B4O7 to neutralize the acid. After washing in PBS – 10% FBS, the cells were resuspended in 200 Al of 0.5% Tween 20 –1% BSA – PBS and incubated with anti-BrdU FITC
(Becton-Dickinson, San Jose, CA, USA) for 30 min at room temperature. The cells were washed with PBS – 10% FBS, and propidium iodide was added to determine the percentage of dead cells. After washing in PBS –10% FBS, the cells were analyzed using a flow cytometer. Table 1a Generation of myeloid DCs from BMMos
I/R 2 d Sham 2 d I/R 14 d Sham 14 d
BMMos ( 105/rat)
DCs (105/rat)
Recovery rate (%)
3647.6 T 341.5 3827.2 T 253.3 4678.0 T 221.5 5156.7 T 172.4
55.3 T 16.2* 90.8 T 22.7 68.3 T 1.2 79.5 T 1.9
1.4 T 0.4* 2.5 T 0.8 1.5 T 0.3 1.6 T 0.4
DCs: dendritic cells; BMMos: bone marrow monocytes. Recovery rate = (DCs BMMos) 100%. * p < 0.05 as compared with the sham 2 d group.
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Measurement of IL-12 IL-12 release from myeloid DCs and plasma IL-12 were measured using an ELISA kit (Biosource Inc., Camarillo, CA, USA) according to the manufacturer’s protocol. Immunohistochemistry of dendritic cells in kidney All rats were perfused via the left cardiac ventricle with about 300 ml of 0.002% heparin sodium in 0.9% sodium
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chloride solution. The kidneys were removed and fixed by immersion in 4% paraformaldehyde (PFA)/PBS at 4 -C overnight, transferred to 30% sucrose/PBS at 4 -C overnight, embedded in Tissue-Tek\ O.C.T compound (Miles Inc., Elkhart, IN, USA), and stored at 80 -C until ready for cryostat sectioning. Sections (5 Am thick) mounted on slides were incubated with mouse monoclonal antibodies specific to CD11c+, CD80, or CD86 (Serotec, Oxford, England) and then with biotin-conjugated goat anti-mouse immunoglobulin and horseradish peroxidase (HRP)-conjugated streptavidin
Fig. 3. DCs stained by antibodies to CD11c A) and D), CD80 B) and E), and CD86 C) and F) are shown in the outer medulla of the kidney at 2 d (A – C) but not at 14 d (D – F) of reperfusion.
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Table 1b Generation of PBDCs from PBMos
I/R 2 d Sham 2 d
Table 2b DC IL-12 secretion by BMDCs
PBMos (105/rat)
PBDCs (105/rat)
Recovery rate (%)
619.0 T 61.5 665.8 T 52.9
21.0 T 2.9 13.5 T 2.9
3.9 T 0.3* 2.1 T 0.5
DCs: dendritic cells; PBMo: peripheral blood monocytes. Recovery rate = (DCs PBMo) 100%. * p < 0.05 as compared with the sham 2 d group.
IL-12 (pg/ml) I/R 2 d Sham 2 d I/R 14 d Sham 14 d
105.1 T 32.1* 217.4 T 51.3 174.2 T 99.6 153.6 T 81.6
IL-12 (pg/105 DCs) 34.6T6.5 38.2 T 12.7 29.1 T17.3 37.6 T 19.4
* p < 0.05 as compared with the sham 2d group.
(DAKO, Glostrup, Denmark). Aminoethylcarbazole was used as chromogenic substrate, and the presence of a red precipitate was identified as positive staining.
differentiation markers on BMMo-derived DCs (Fig. 2C) was not significantly different at both 2 and 14 d. Dendritic cell infiltration of the kidney
Statistical analysis Data are expressed as the mean T SE. All comparisons were made by analysis of variance followed by Dunnett’s t-test using the Statview software package for the Macintosh computer (Abacus Concepts, Berkeley, CA, USA). Results
Immunohistochemical analysis of CD11c, CD80, and CD86 labeling of DCs in kidney sections from I/R injured rats showed DCs are present in the outer medulla at 2 d (Fig. 3A – C) but not 14 d of reperfusion (Fig. 3D – F). DCs were not seen in kidney sections of sham-operated rats at either time. Generation of peripheral blood monocyte-derived DCs at 2 d of reperfusion
Changes in renal function after I/R injury Renal dysfunction, indicated by elevated plasma creatinine levels and decreased calculated creatinine clearance, was observed on d 0 through 4 (peak day 1—0.32 T 0.12 ml/min) of reperfusion in rats subjected to I/R (Gr 1 and Gr 3) compared to sham-operated rats (Gr 2 and Gr 4) (data not shown). Generation of bone marrow monocyte-derived DCs Myeloid DCs generated from BM cells cultured in the presence of rGM-CSF and rIL-4 for six days and in the presence of LPS for one additional day were recovered as a population of nonadherent or loosely adherent clustered and veiled cells (Fig. 1) and recognized by flow cytometry as a population of CD11c-CD80-CD86-Ia-bearing cells (Fig. 2A and B). In preliminary experiments, no significant difference in DC differentiation was observed between d 1 and 5 of reperfusion (data not shown). However, the number of mature BMMo-derived DCs was significantly lower at 2 d after reperfusion in I/R rats than in sham-operated rats (Gr 1 vs. Gr 2), whereas this number was not significantly different at 14 d (Gr 3 vs. Gr 4) (Table 1a) and the per-cell expression of Table 2a Mixed lymphocyte response of peripheral blood (PB) and splenic Tlymphocytes to stimulator bone marrow dendritic cells (BMDCs) BMDCs from rats treated with:
Control
PB T-cells
Control
Splenic T-cells
I/R 2 d Sham 2 d I/R 14 d Sham 14 d
9.6 T 1.6 9.6 T 1.6 4.5 T 1.0 4.5 T 1.0
32.5 T 4.2 31.5 T 4.6 26.0 T 6.9 21.9 T 6.8
3.1 T 0.4 3.1 T 0.4 4.0 T 1.0 4.0 T 1.0
20.3 T 4.4 22.4 T 4.1 25.3 T 9.4 20.7 T 6.2
The results are expressed as percent of BrdU (5-bromo-2-deoxyuridine)positive cells and means T SEM for N = 12. No significant differences between I/R- and sham-treated animals are shown.
Because BMMo differentiation to DCs decreased at 2 d at the same time that DCs increased in the outer medulla of the kidney, we thought that PBMos might also be responders at 2 d of reperfusion to ischemic injury. Hence, PBMo-derived DCs were counted and found to be significantly higher at 2 d (Table 1b). Mixed lymphocyte reaction stimulated by BMMo-derived DCs was unaffected by I/R injury Whether the source of T cells was peripheral blood or spleen (Table 2a), no significant differences in MLR proliferation were found between I/R rats and sham-operated rats. IL-12 production of BMMo-derived DCs was unaffected by I/R injury Though myeloid DCs from I/R rats produced considerably less IL-12 by 2 d of reperfusion than myeloid DCs from shamoperated rats, the difference reflected merely the decrease in cell numbers. There was no significant difference in the per 105 cell release of IL12 between the groups (Gr 1 vs. Gr 2 and Gr 3 vs. Gr 4; Table 2b). In plasma, IL-12 levels were undetectable in all groups (detection level, 0 –500 pg/ml). Discussion Our study suggests that I/R injury 1) significantly decreases the number of myeloid DCs differentiated from BMMos at 2 d but not at 14 d of reperfusion 2) has no effect on functional activity (i.e., no change in DC-stimulated MLR and in IL-12 synthesis of individual DCs), and 3) increases the number of DCs in the kidney at 2 d. These changes may reflect inhibition of DC differentiation from BMMos and either stimulation of
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DC differentiation from Mos in the kidney or migration of DCs to the kidney during the acute phase of I/R injury. I/R injury was also found to increase the number of DCs differentiated from peripheral blood monocytes (PBMos) at 2 d. The difference between the differentiability of monocytes in BM and monocytes in PB suggests that these cells represent two distinct subpopulations. In support of this possibility, Sunderkotter et al. have demonstrated in mice that immature Ly6Chigh monocytes (phenotypically BMMos) immigrate from the BM to the circulation, and that during inflammation they predominate for > 2 d in the circulation where they gradually mature to Ly6Clow monocytes (phenotypically PBMos). Under steady state conditions, the predominant monocytes in the circulation were the more mature Ly6Clow cells, whereas under inflammatory conditions, the most abundant were immature Ly6Chigh (the phenotype in bone marrow) and less mature Ly6Cmedium cells (Sunderkotter et al., 2004). These findings imply that the primary distinction between monocyte subsets is their maturation state and that environmental factors such as inflammatory conditions determine subset predominance. Similarly, another study found that maturation state and therefore DC subset is influenced by environmental factors: the presence of CD40 ligand (which accelerates maturation of DCs) and phagocytosis of bacteria and yeasts (which favors development of less mature DCs) (Rosenzwajg et al., 2002). Thus, microenvironmental influences appeared to direct monocyte maturation and subset development. Our study also showed I/R injury resulted in DC infiltration of the outer medulla of the kidney after 2 d but not 14 d of reperfusion. We can only speculate that the transience of DC appearance reflects either DC entry followed by DC exodus from the kidney or further maturation of mature DCs, e.g., into endothelial-like cells (Havemann et al., 2003). Analogously, Kennedy and Abkowitz found that the more mature monocyte subpopulation of their monocyte transplant landed in various organs 3 d after transplantation and that by 7 d, these cells had differentiated into macrophages in situ. At 3 d, cells were found as clusters and individual cells. By 7 and 14 d, the clusters had increased in size and the cells expressed the macrophage antigen F4/80, suggesting that further replication and differentiation had occurred (Kennedy and Abkowitz, 1998). Our results are consistent with this idea that bone marrow or peripheral blood DCs or DC progenitors migrate to kidney via peripheral blood following I/R injury. Also, Sunderkotter et al. observed in mice experimentally depleted of monocytes the presence of repopulating macrophages and DCs in peripheral organs (particularly spleen and liver) 4 d after monocyte depletion (Sunderkotter et al., 2004). The functional role of DCs that settle in the renal medulla as a result of I/R injury is unclear. This will require further investigation. As for the fate of these DCs, our study found that the DC response to I/R injury (decreased differentiation from BMMos, increased differentiation from PBMos, and infiltration of the renal outer medulla, which is the area susceptible to I/R injury [Brezis et al., 1984]) occurred only during the acute phase (d 2) of renal I/R injury. This finding suggests 1) that DC differentiation from BMMos may shift to PBMos, then DCs
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may migrate from PB to the kidney and 2) that this shift and migration are stress-induced. Because our study focused on differentiation and did not track the migration of DCs, we do not know whether the decreased DCs in BM is due to migration to PB and then to kidney. Penfield et al. have shown that injury to the kidney during transplantation is sufficient to recruit recipient MHC class II leukocytes (some of which are DCs) into the transplant (Penfield et al., 1999). Similarly, Loi et al. showed, in a mouse model, that liver I/R injury itself induces DC maturation, migration, and preferential production of inhibitory cytokines (Loi et al., 2004). However, they failed to show whether the DCs in the liver parenchyma were recruited in situ or were derived from circulating precursors. Because PBDCs, BMDCs, and DCs infiltrating the renal medulla may be different subpopulations and because these cells may interact with and stimulate different T cell populations, comparison of these cells is uncertain and the problem requires further investigation. Two aspects of the MLR technique used in this study should be mentioned because they may be regarded as limitations. First, the technique was modified (BrdU [a nonradioactive analogue of thymidine] rather than 3H-thymidine was measured (Gratzner, 1982)), and second, a mixed T cell population rather than bead-purified T cells were added. In summary, the upshift in PBMo differentiation to DCs, downshift in BMMo differentiation to DCs, and infiltration of the kidney at 2 d of reperfusion may be part of an important pathophysiologic acute stress response to renal I/R injury. Acknowledgments We thank Gabriel Cheng, Ming-Ling Hsu, Hui-Ju Shieh, Hung-Jen Liao and Ching-Pin Lin for their technical help and writing assistance. This work was supported by research grants from the National Science Council (NSC 93-2413-H-195-001) and Mackay Memorial Hospital Medical Research Fund (MMHE93006 and MMH9372). References Bonventre, J.V., Weinberg, J.M., 2003. Recent advances in the pathophysiology of ischemic acute renal failure. Journal of the American Society of Nephrology 4, 2199 – 2210. Bonsnes, R.M., Taussky, H.M., 1945. The colorimetric determination of creatinine by Jaffe’s reaction. Journal of Biological Chemistry 158, 581 – 591. Brezis, M., Rosen, S., Silva, P., Epstein, F.H., 1984. Renal ischemia: a new perspective. Kidney International 26 (4), 375 – 383. De Vecchi, E., Lubatti, L., Beretta, C., Ferrero, S., Rinaldi, P., Galli Kienle, M., Trazzi, R., Paroni, R., 1998. Protection from renal ischemia – reperfusion injury by the 2-methylaminochroman U83836E. Kidney International 54 (3), 857 – 863. Gratzner, H.G., 1982. Monoclonal antibody to 5-bromo- and 5-iododeoxyuridine: a new reagent for detection of DNA replication. Science 218, 474 – 475. Havemann, K., Pujol, B.F., Adamkiewicz, J., 2003. In vitro transformation of monocytes and dendritic cells into endothelial like cells. Advances in Experimental Medicine and Biology 522, 47 – 57. Kennedy, D.W., Abkowitz, J.L., 1998. Mature monocytic cells enter tissues and engraft. Proceedings of the National Academy of Sciences of the United States of America 95 (25), 14944 – 14949.
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C.-J. Wu et al. / Life Sciences 78 (2006) 1121 – 1128
Lieberthal, W., Nigam, S.K., 1998. Acute renal failure. I. Relative importance of proximal vs. distal tubular injury. American Journal of Physiology 275, F623 – F631. Loi, P., Paulart, F., Pajak, B., Nagy, N., Salmon, I., Moser, M., Goldman, M., Flamand, V., 2004. The fate of dendritic cells in a mouse model of liver ischemia/reperfusion injury. Transplantation Proceedings 36 (5), 1275 – 1279. Penfield, J.G., Wang, Y., Li, S., Kielar, M.A., Sicher, S.C., Jeyarajah, D.R., Lu, C.Y., 1999. Transplant surgery injury recruits recipient MHC class II-positive leukocytes into the kidney. Kidney International 56 (5), 1759 – 1769. Rissoan, M.C., Soumelis, V., Kadowaki, N., Grouard, G., Briere, F., de Waal Malefyt, R., Liu, Y.J., 1999. Reciprocal control of T helper cell and dendritic cell differentiation. Science 283 (5405), 1183 – 1186. Rosenzwajg, M., Jourquin, F., Tailleux, L., Gluckman, J.C., 2002. CD40 ligation and phagocytosis differently affect the differentiation of monocytes into dendritic cells. Journal of Leukocyte Biology 72 (6), 1180 – 1189. Rutault, K., Alderman, C., Chain, B.M., Katz, D.R., 1999. Reactive oxygen species activate human peripheral blood dendritic cells. Free Radical Biology & Medicine 26 (1 – 2), 232 – 238.
Sallusto, F., Lanzavecchia, A., 1994. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. Journal of Experimental Medicine 179, 1109 – 1118. Sunderkotter, C., Nikolic, T., Dillon, M.J., Van Rooijen, N., Stehling, M., Drevets, D.A., Leenen, P.J., 2004. Subpopulations of mouse blood monocytes differ in maturation stage and inflammatory response. Journal of Immunology 172 (7), 4410 – 4417. Talmor, M., Mirza, A., Turley, S., Mellman, I., Hoffman, L.A., Steinman, R.M., 1998. Generation or large numbers of immature and mature dendritic cells from rat bone marrow cultures. European Journal of Immunology 28 (3), 811 – 817. Verhasselt, V., Goldman, M., Willems, F., 1998. Oxidative stress up-regulates IL-8 and TNF-alpha synthesis by human dendritic cells. European Journal of Immunology 28, 3886 – 3890. Vieira, P.L., de Jong, E.C., Wierenga, E.A., Kapsenberg, M.L., Kalinski, P., 2000. Development of Th1-inducing capacity in myeloid dendritic cells requires environmental instruction. Journal of Immunology 164 (9), 4507 – 4512.