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Absence of angiotensin II type 1 receptor in bone marrow–derived cells is detrimental in the evolution of renal fibrosis
Masashi Nishida,1 Hidehiko Fujinaka,1 Taiji Matsusaka,1,2 James Price,3 Valentina Kon,1 Agnes B. Fogo,1,3,4 Jeffrey M. Davidson,3 MacRae F. Linton,4 Sergio Fazio,4 Toshio Homma,2 Hiroaki Yoshida,1,4 and Iekuni Ichikawa1,2,4
1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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1Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee, USA2 Department of Pediatrics, Tokai University School of Medicine, Isehara, Japan3 Department of Pathology, and4 Department of Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA
Address correspondence to: Iekuni Ichikawa, MCN C4204, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2584, USA. Phone: (615) 322-7931; Fax: (615) 322-7929; E-mail: ichikawamd@aol.com.
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Published December 15, 2002 - More info
J Clin Invest. 2002;110(12):1859–1868. https://doi.org/10.1172/JCI15045.
© 2002 The American Society for Clinical Investigation
Received: January 14, 2002; Accepted: October 1, 2002
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Methods
Mice. Agtr1–/– donor mice used in the present study were generated in our laboratory (24) and had been backcrossed more than 10 times with C57BL/6 mice (The Jackson Laboratory, Bar Harbor, Maine, USA). Thus, all mice used in the study (both the genetically manipulated donors and the wild-type recipient mice) were on the C57BL/6 background. Rodents carry two subtypes of the Agtr1 receptor — namely, type 1a and type 1b (25). Previous data have demonstrated that the angiotensin II type 1a receptor is by far the predominant isoform in mice (26). We therefore used the angiotensin II type 1a receptor null mutant mice, designated as Agtr1–/– mice, in the present study. The genotype of these mice was determined by Southern blot analysis on DNA isolated from tail biopsies as previously described (24). Male C57BL/6 mice were purchased from The Jackson Laboratory.
Bone marrow reconstitution and unilateral ureteral obstruction. Bone marrow transplantation was performed as previously described (22, 23). Three-month-old male recipient mice were lethally irradiated with 9 Gy using a cesium gamma source. Bone marrow cells were harvested from the femurs and tibias of donor mice by flushing with RPMI-1640 media (Gibco BRL, Gaithersburg, Maryland, USA), and recipients received 5 × 106 bone marrow cells per mouse in 0.2 ml of media by tail-vein injection. To verify the reconstitution of bone marrow after transplantation, the bone marrow and peritoneal macrophages were collected at the time of sacrifice, genomic DNA was extracted from these cells, and the genotype was determined by Southern blot analysis as we have previously described (24).
Eight weeks after bone marrow transplantation, the transplanted mice were subjected to a complete unilateral ureteral obstruction (UUO) as previously described (27). Briefly, under sodium pentobarbital anesthesia, the middle portion of the left ureter was ligated and cut between the two ligated points. At 5 and 14 days after surgery, the mice were sacrificed, and the obstructed kidneys were harvested and subjected to the studies described below. The experimental protocols were performed according to the regulations of the Vanderbilt University Animal Care Committee.
Histological study. For histological examinations, kidneys were fixed with 4% buffered paraformaldehyde for 6 hours, embedded in paraffin, and sectioned transversely with a thickness of 4 μm. A standard point-counting method was used to quantitate the collagen fractional volume in the renal cortical interstitium on Masson trichrome–stained sections (magnification, ×400), as described in our previous studies (27). The index of interstitial collagen fractional volume was defined as the number of trichrome-positive points in every 1000 points evaluated. For immunohistochemistry, monoclonal rat anti-mouse F4/80 (Selotec, Oxford, United Kingdom) and anti-mouse CD3-ε (clone 48-2B; Santa Cruz Biotechnologies Inc., Santa Cruz, California, USA) were used to detect infiltrat-ing macrophages and T cells, respectively (27, 28). α-Smooth muscle actin (α-SMA) was stained by mouse monoclonal anti-human α-SMA (clone 1A4; DAKO, Carpinteria, California, USA) (27, 28). Vascular endothelial cells were stained with a monoclonal rat antibody to a cell membrane antigen present on endothelial cells, CD31 (clone MEC 13.3, 1:50 dilution; Pharmingen, San Diego, California, USA) (29). After exposure to 0.1% trypsin (DAKO) for 10–15 minutes and quenching with 3% hydrogen peroxide in methanol, sections were incubated with rat anti-mouse CD31 for 1 hour at room temperature, followed by standard ABC immunostaining.
Macrophage infiltration was determined by enumerating F4/80-positive cells within the cortical interstitium in 10 randomly selected cortical fields under magnification (×400), and the numbers were averaged for each field (27). The density of CD31-positive peritubular capillary was quantified as the peritubular capillary rarefaction index as previously described (30). Briefly, the squares in a 10 × 10 grid that showed no capillaries were counted in 10 sequential fields under magnification (×200) and averaged. The expression of α-SMA in the cortical interstitial area was assessed semiquantitatively by immunoreactivity for α-SMA as previously described (28).
Collagen content. The collagen content of renal cortex was calculated from the concentrations of hydroxy-proline and proline, which were determined through measurement of their phenylisothiocyanate derivatives by reverse-phase high-performance liquid chromatography (RP-HPLC) as previously described (31).
Northern blot analyses for TGF-β1, α1(I) collagen, α1(III) collagen, matrix metalloproteinase 2, and tissue inhibitor of metalloproteinase 1 mRNA. Total RNA was extracted by the RNA STAT-60 method (Tel-Test Inc., Friendswood, Texas, USA), electrophoresed, transferred to a nylon membrane (Hybond-N+; Amersham Life Science Inc., Arlington Heights, Illinois, USA), and hybridized with a 32P-labeled probe. The cDNA probes for mouse TGF-β1, α1(I) collagen, α1(III) collagen, matrix metalloproteinase-2 (MMP-2), and tissue inhibitor of metalloproteinase 1 (TIMP-1) were used as described previously (27, 28). The hybridization signal of each mRNA was detected by autoradiography, scanned with a JX-330M scanner (Sharp Co., Tokyo, Japan), and semiquantitated using the National Institutes of Health image program.
Hematologic analysis. Total leukocyte, red blood cell, and platelet counts were determined on peripheral blood collected by retro-orbital venous plexus puncture using an ADVIA 120 automated counter (Bayer, Tarrytown, New York, USA), and the manual 100-cell leukocyte differential counting was performed on May-Grüwald/Giemsa–stained smears.
Determination of bone marrow macrophage progenitors. Macrophage progenitors present in the bone marrow were determined as granulocyte or macrophage colony-forming unit by semisolid agar culture (32). Bone marrow cells were harvested by flushing the tibia with RPMI-1640 media and were cultured for 10 days at a density of 50,000 cells per milliliter in media consisting of RPMI-1640, 0.3% agar, 10% FBS, and recombinant mouse colony stimulating factor 1 (CSF-1) at concentrations indicated in Figure 6. Macrophage and granulocyte/macrophage colonies containing more than 50 cells were morphologically identified in triplicate cultures, counted, and expressed as colony-forming units per tibia.
Figure 6Macrophage progenitors in bone marrow. Bone marrow cells isolated from either Agtr1+/+ mice or Agtr1–/– mice were cultured for 10 days in semisolid agar culture at a density of 50,000 cells per milliliter in the presence of indicated concentrations of CSF-1. Colonies of macrophages or granulocyte/macrophages were counted (a), and the total number of macrophage progenitors per tibia was determined (b). *P < 0.05.
Determination of macrophage migrating activity. Peritoneal macrophages were collected from mice 4 days after peritoneal injection of 3% thioglycollate. Chemotaxis was performed in a 96-well microchemotaxis chamber in which the upper compartment was separated from the lower compartment by a single uncoated polycarbonate filter (5-μm pore diameter; Neuroprobe, Cabin John, Maryland, USA). Peritoneal macrophages were suspended in DMEM without FBS and added to the upper compartment of the chemotaxis chamber at a density of 1.2 × 106 cells per well. Chemoattractants were added to the lower compartment, and chambers containing cells and chemoattractants were incubated at 37°C for 2.5 hours in an atmosphere of 95% air, 5% CO2, and 100% humidity. After incubation, cells remaining on the upper surface of the filter were removed mechanically. Filters were then fixed in methanol and stained with 1% crystal violet. Migrated cells adhering to the lower surface of the membrane were counted manually under the microscope with an oil-immersion lens (×400). Triplicate wells were used for each experimental condition, and 10 fields were examined for each well.
Measurement of Fcγ receptor–mediated oxidative burst. Fcγ receptor–mediated oxidant production in peritoneal macrophages was measured using the FcOxyBurst reagent (Molecular Probes, Eugene, Oregon, USA) according to the manufacturer’s instructions. Briefly, peritoneal macrophages harvested from mice 4 days after peritoneal injection of 3% thioglycollate were suspended at 2 × 106 cells per milliliter in PBS supplemented with 1g/l glucose and then stimulated with FcOxyBurst at a final concentration of 140 μg/ml. Flow cytometric analysis was performed with a FACScan flow cytometer (Becton Dickinson, Franklin Lakes, New Jersey, USA) gating to select F4/80-positive macrophages. Twenty thousand 7-amino-actinomycin D–negative (7-AAD–negative), living cells were obtained for the analysis.
In vivo and in vitro assays of phagocytic activity of macrophages. Phagocytic activity of macrophages in vivo was examined using previously described methods (33, 34). Four days after thioglycollate injection, 2.5 ml of a 0.2 % suspension of Fluoresbrite fluorescent microspheres (0.75 YG; Polyscience, Warrington, Pennsylvania, USA) was injected into the peritoneal cavity. After 1 hour of incubation, peritoneal fluid containing cells and fluorescent beads was collected. The macrophages with fluorescent beads were counted in accordance with the amount of ingested beads (34), and the nonphagocytosed free beads were also quantified by flow cytometry as a percentage of the total number of beads (33) on a FACScan flow cytometer using CellQuest software (Becton Dickinson) for data acquisition and Winlist software (Verity Software House, Topsham, Maine, USA) for data analysis. Twenty thousand 7-AAD–negative, living cells were obtained for the analysis. The flow cytometric analysis was performed gating to select F4/80-positive macrophages, and the fluorescence of extracellular beads was quenched using crystal violet according to the manufacturer’s protocol. To determine the role of Agtr1 in vivo, losartan was added to the drinking water (200 mg/l) 18 hours before the assay, and a bolus of 1.0 mg was also injected into the peritoneal cavity with fluorescent beads.
For the in vitro phagocytic assay, peritoneal macrophages were collected from mice 4 days after peritoneal injection of 3% thioglycollate, and approximately 2 × 106 peritoneal macrophages on 24-well plates in DMEM with 10% FBS were used (33, 35). One hundred microliters of 0.26% latex solution of Fluoresbrite fluorescent microspheres (0.75 YG; Polyscience) was added to each well and incubated for 30 minutes at 37°C. Nonadherent latex beads were removed by washing, and the cells were incubated for another 30 minutes to allow for complete phagocytosis. Cells were then harvested by treatment with PBS containing 5 mM EDTA and washed vigorously three times with PBS to remove nonphagocytosed beads. The amount of phagocytosed beads was measured by flow cytometry as described above. In some experiments, either the Agtr1 antagonist losartan or angiotensin II (Bachem, Torrance, California, USA) was added to the final concentration of 10–6 M 18 hours or 3 hours before the phagocytosis assay, respectively. The amount of nonspecifically bound beads was determined by incubating macrophages with fluorescent beads on ice.
Statistical analysis. Data are presented as means ± SEM. Statistical analysis was performed by ANOVA, and statistical significance was defined as P < 0.05.
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Results
Reconstitution of Agtr1+/+ mice with Agtr1–/– bone marrow. Resident macrophages of the recipient mice were removed from the peritoneal cavity at the time of sacrifice and genotyped (Figure 1). Uniformly, only the 5-kb band was observed in the recipient mice, demonstrating that Agtr1+/+ mice were successfully reconstituted with Agtr–/– marrow, and macrophages were primarily from the donor (lane 4 in Figure 1). This finding is in good agreement with our previous bone marrow reconstitutions (22).
Figure 1Southern blot genotype analysis of macrophages isolated from mice transplanted with either Agtr1+/+ or Agtr1–/– bone marrow. Lanes 1–3 show DNA isolated from tail biopsy specimens of Agtr1–/– mice (Lane 1), Agtr1+/– mice (Lane 2), and Agtr1+/+ mice (Lane 3). Lanes 4 and 5 show DNA isolated from peritoneal macrophages of wild-type mice reconstituted with Agtr1–/– marrow (Lane 4) and wild-type mice with Agtr1+/+ marrow (Lane 5). The DNA in lane 4 represents pooled macrophage DNA from 10 mice reconstituted with Agtr1–/– marrow.
Effect of Agtr1–/– bone marrow transplantation on interstitial macrophage infiltration in the obstructed kidneys. Macrophage accumulation in the kidney was examined in wild-type mice with Agtr1–/– marrow and wild-type mice with Agtr1+/+ marrow at day 5 (Agtr1–/– marrow, n = 9; Agtr1+/+ marrow, n = 10) and day 14 (Agtr1–/– marrow, n = 8; Agtr1+/+ marrow, n = 10) after surgical creation of UUO. At day 5, there was no appreciable difference in the number of interstitial macrophages between the two groups (17.3 ± 2.5 vs. 19.7 ± 3.0, not significant) (Figure 2b). At 14 days after UUO, mice carrying transplanted Agtr1–/– marrow showed significantly lower numbers of F4/80-positive interstitial macrophages than those carrying Agtr1+/+ marrow (27.6 ± 1.5 vs. 41.8 ± 2.5, P < 0.001) (Figure 2, a and b). Staining for CD3 revealed that there were only a few T cells in the interstitium in both groups at either day 5 or day 14 after UUO. No significant difference was observed between the groups (data not shown).
Figure 2Macrophage infiltration after UUO in wild-type mice with either Agtr1–/– marrow or Agtr1+/+ marrow. (a) Immunohistochemical demonstration of macrophages with anti-F4/80 antibody in mice with Agtr1–/– marrow (left) and mice with Agtr1+/+ marrow (right) at day 14 after UUO. Magnification, ×400. (b) Macrophages (F4/80-positive cells) infiltrating to the interstitium at day 5 (left) and day 14 (right) after UUO. Black bars represent mice with Agtr1–/– marrow; white bars represent mice with Agtr1+/+ marrow. Data are expressed as means ± SEM. *P < 0.001 for mice with Agtr1–/– marrow versus mice with Agtr1+/+ marrow. NS, not significant.
Effect of Agtr1–/– bone marrow transplantation on renal fibrosis in the obstructed kidneys. The degree of fibrosis was compared between wild-type mice with Agtr1–/– marrow or Agtr1+/+ marrow at day 5 and day 14 after UUO (Figure 3a). At day 5 after UUO, there was no difference in interstitial fibrosis (44 ± 8 vs. 42 ± 6 points per 1000 points, not significant) (Figure 4a). However, at 14 days after UUO, mice carrying Agtr1–/– marrow (n = 8) showed significantly more severe interstitial fibrosis than those carrying Agtr1+/+ marrow (n = 10; 139 ± 9 vs. 93 ± 9 points per 1000 points, P < 0.005) (Figure 4a). The enhanced fibrosis seen in mice carrying Agtr1–/– marrow was confirmed independently by a biochemical determination of collagen content. Thus, collagen content in renal tissue assessed by RP-HPLC was also significantly higher in mice with Agtr1–/– marrow than in mice with Agtr1+/+ marrow (11.6% ± 1.1% vs. 7.7% ± 0.5% collagen, P < 0.005) (Figure 4b).
Figure 3Histological assessment of interstitial fibrosis and peritubular capillary density and detection of myofibroblasts in the obstructed kidneys of wild-type mice with either Agtr1–/– marrow or Agtr1+/+ marrow. Masson trichrome staining of the renal cortex at day 14 after UUO (a), immunohistochemical assessment of peritubular capillary density by immunostaining with anti-CD31 antibody at day 5 after UUO (b), and detection of myofibroblasts by immunostaining with anti-αSMA antibody at day14 after UUO (c) in mice with Agtr1–/– marrow (left) and mice with Agtr1+/+ marrow (right). Magnification, ×400.
Figure 4Quantitative analyses of interstitial collagens after UUO in wild-type mice with either Agtr1–/– marrow or Agtr1+/+ marrow. (a) Interstitial collagen index assessed by point-counting method on Masson trichrome–stained sections at day 5 (left) and day 14 (right) after UUO. (b) Collagen content as percentage of total protein in kidney at day 14 after UUO. Black bars represent mice with Agtr1–/– marrow; white bars represent mice with Agtr1+/+ marrow. Data are expressed as means ± SEM.*P < 0.005 for mice with Agtr1–/– marrow versus mice with Agtr1+/+ marrow.
Using CD31 staining, we also evaluated peritubular capillary density. The new generation of peritubular capillaries is believed to be promoted by macrophage-derived bFGF in early stages of UUO (36). No significant difference in the peritubular capillary density was seen between the two groups 5 days after UUO (rarefaction index, 4.3% ± 0.5% vs. 4.9% ± 0.7%, not significant) (Figure 3b). Transdifferentiation of tubular epithelial cells to myofibroblasts has been implicated as another component of the fibrogenetic processes in UUO (37). Cells with α-SMA–positive staining were located in the renal interstitium and vessel walls (Figure 3c). We found no significant difference in the immunoreactivity of α-SMA between the two groups 14 days after UUO (α-SMA score, 2.66 ± 0.06 vs. 2.44 ± 0.10, not significant).
mRNA expressions for molecules involved in the accumulation of ECM. In parallel with the pattern of interstitial fibrosis, expression of TGF-β1 mRNA was significantly higher in mice with Agtr1–/– marrow than in mice with Agtr1+/+ marrow at day 14 after UUO (the ratio of optical density to that of GAPDH mRNA, 2.08 ± 0.23 vs. 0.59 ± 0.08, P < 0.001). TGF-β1 mRNA expression was similar at day 5 (1.07 ± 0.12 vs. 1.19 ± 0.09, not significant) (Figure 5a). In mice with Agtr1+/+ marrow, TGF-β1 mRNA expression was not significantly different between day 5 and day 14. In contrast, in mice with Agtr1–/– marrow, TGF-β1 mRNA expression was significantly increased by day 14 from the level seen on day 5 (Figure 5a).
Figure 5mRNA expressions of molecules involved in the accumulation of ECM. (a) Northern blot analyses of TGF-β1 expressions at day 5 (left) and day 14 (right) after UUO in wild-type mice with Agtr1–/– marrow and wild-type mice with Agtr1+/+ marrow. (b and c) Northern blot analyses of α1(I) collagen, MMP-2, and TIMP-1 at day 5 (b) and day 14 (c) after UUO. Lane 1 in (b) and (c) shows control contralateral nonobstructed kidney, lane 2 shows mice with Agtr1–/– marrow, and lane 3 shows mice with Agtr1+/+ marrow.
Figure 5 (b and c) summarizes mRNA expression for molecules involved in accumulation as well as those involved in degradation of ECM. Profibrotic molecules, including α1(I) collagen and α1(III) collagen, were expressed at substantially higher levels in mice with Agtr1–/– marrow than in mice with Agtr1+/+ marrow at day 14 (α1[I] collagen: 1.93 ± 0.39 vs. 0.85 ± 0.14, P < 0.05; α1[III] collagen: 2.79 ± 0.61 vs. 1.25 ± 0.11, P < 0.05). Likewise, the expression of MMP-2 mRNA was higher in mice with Agtr1–/– marrow than those with Agtr1+/+ marrow (2.40 ± 0.43 vs. 0.74 ± 0.24, P < 0.01), whereas no significant difference was ob-served in the expression of TIMP-1 mRNA at day 14 after UUO (1.20 ± 0.36 vs. 1.83 ± 0.20) (Figure 5c). At day 5 after UUO, no significant difference was observed between mice with Agtr1–/– marrow or Agtr1+/+ marrow in α1(I) collagen mRNA (1.27 ± 0.14 vs. 1.38 ± 0.13), α1(III) collagen mRNA (0.60 ± 0.08 vs. 0.49 ± 0.09), MMP-2 mRNA (0.98 ± 0.11 vs. 1.00 ± 0.12), or TIMP-1 mRNA (1.57 ± 0.16 vs. 1.71 ± 0.18) (Figure 5b).
Peripheral monocytes and bone marrow macrophage progenitors. Interstitial macrophages are the major infiltrating cells in UUO and are believed to be an active element of the interstitial fibrosis. Therefore, to unveil the mechanisms underlying the reduced macrophage infiltration and the enhanced fibrosis seen in wild-type mice with Agtr1–/– marrow, we examined a few features of macrophages. The number of circulating hema-topoietic cells was examined in wild-type mice with Agtr1–/– marrow or Agtr1+/+ marrow 8 weeks after bone marrow transplantation (Table 1). Leukocytes were significantly less abundant in mice with Agtr1–/– marrow than in those with Agtr1+/+ marrow. Although the number of monocytes in mice with Agtr1–/– marrow was significantly lower than in those with Agtr1+/+ marrow, the monocyte/white blood cell ratio was similar between the two groups of mice. Thus, the difference is not unique to monocytes.
Total bone marrow cells per tibia were not significantly different between mice with Agtr1+/+ or Agtr1–/– marrow (22.3 ± 4.1 × 106 cells per tibia vs. 17.6 ± 3.8 × 106 cells per tibia, not significant). When 50,000 marrow cells were cultured for 10 days in semisolid agar culture in the presence of increasing concentrations of cerebrospinal fluid 1 (CSF-1), growth of macrophage progenitor colonies reached a plateau at 5 ng/ml of CSF-1 for both Agtr1+/+ and Agtr1–/– marrow (Figure 6a). Thus, this assay showed that mice with Agtr1–/– marrow had significantly less macrophage progenitors than those with Agtr1+/+ marrow (18.0 ± 2.8 × 103 cells per tibia vs. 25.5 ± 3.2 × 103 cells per tibia, P < 0.05) (Figure 6b).
Functions of macrophages. To explore further the mechanisms responsible for the enhanced fibrosis in mice with Agtr1–/– marrow, we next examined macrophage functions. Monocyte chemotactic protein 1 (MCP-1) has been shown to be the major chemotactic factor responsible for macrophage infiltration in UUO (38), and chemotactic activity of angiotensin II has been demonstrated for human monocytes (39). On stimulation with 0.1 μg/ml MCP-1, Agtr1+/+ macrophages exhibited a maximal 1.5-fold increase in chemotactic response from 124 ± 6 cells per field to 186 ± 41 cells per field (left panel in Figure 7a). In response to 0.1 μg/ml MCP-1, a similar 1.5-fold increase was observed in the chemotactic response of Agtr1–/– macrophages from 34 ± 7 cells per field to 51 ± 11 cells per field, indicating that although the baseline motility of Agtr1–/– macrophages is lower than that of Agtr1+/+ macro-phages, macrophages of either genotype have indistinguishable chemotactic responses to MCP-1. Addition of 1 × 10–6 M angiotensin II, either alone or with 0.1 μg/ml MCP-1, was without effect on the motility of Agtr1+/+ macrophages (right panel in Figure 7a).
Figure 7Migrating activity and oxidative burst of Agtr1–/– and Agtr1+/+ macrophages. (a) Migrating activity of isolated peritoneal macrophages was determined by using chemotaxis chambers. In the left panel, Agtr1–/– or Agtr1+/+ macrophages were incubated with either vehicle (black bars) or 0.1 μg/ml MCP-1(white bars). In the right panel, Agtr1+/+ macrophages were exposed to vehicle (white bar), 1 × 10–6 M angiotensin II alone (gray bar), or 0.1 μg/ml MCP-1 and 1 × 10–6 M angiotensin II (black bar). Migrated cells were counted and are shown as the number of migrated cells per field. *P < 0.05 for vehicle versus MCP-1. (b) Macrophages of the Agtr1–/– (black bar) or the Agtr1+/+ (white bar) genotype were examined for Fcγ receptor–mediated oxidative burst. Oxidative burst was detected as fluorescence emitted by macrophages and is indicated in arbitrary units. Ang II, angiotensin II; MCP-1, monocyte chemotactic protein-1.
Reactive oxygen species are potent destructive effector molecules produced by macrophages and are thought to be involved in the renal damage occurring in UUO (40). On stimulation through the Fcγ receptor, Agtr1+/+ and Agtr1–/– macrophages showed similar oxidative burst (mean fluorescence intensity of total macrophages after 15 minutes of incubation, 35.8 ± 0.9 vs. 38.1 ± 0.9, not significant) (Figure 7b), suggesting that together with reduced macrophage infiltration, macrophage-mediated oxidative injury would be less significant in mice with Agtr1–/– macrophages than in those with Agtr1+/+ macrophages.
Active removal of ECM has been suggested as a critical element that maintains the dynamic balance of the highly regulated wound-healing process, and macrophages are thought to participate in ECM removal through phagocytosis (13). To examine the phagocytic capability of macrophages in vivo, we injected fluorescent beads into the peritoneal cavity after inducing macrophages with thioglycollate. Analysis of the peritoneal fluid obtained 1 hour later showed that both the percentage of macrophages with phagocytosed beads and the mean fluorescence intensity were greater in Agtr1+/+ mice than in Agtr1–/– mice (85.6% ± 1.2% vs. 49.8% ± 1.4 % phagocytic macro-phages, P < 0.001; mean fluorescence intensity of total macrophages, 208.6 ± 12.7 vs. 109.4 ± 8.3, P < 0.005) (Figure 8a). Similarly, fewer fluorescent beads were free (nonphagocytosed) in Agtr1+/+ than in Agtr1–/– mice (remaining free beads, 56.8% ± 6.8% vs. 81.8% ± 3.5 %, P < 0.005; duplicated experiments, n = 6 per group). The effect of losartan on the phagocytic capability of Agtr1+/+ macrophages was also examined in vivo. The losartan treatment caused a significant decrease in the phagocytic capability of Agtr1+/+ macrophages in vivo (85.6% ± 1.2% vs. 47.3% ± 2.2% phagocytic macro-phages, P < 0.001; mean fluorescence intensity of total macrophages, 208.6 ± 12.7 vs. 99.1 ± 5.1, P < 0.005; remaining free beads, 56.8% ± 6.8% vs. 82.9% ± 4.8 %, P < 0.05) (Figure 8a). Thus, the phagocytic capability of Agtr1–/– macrophages is reduced in vivo. Furthermore, the Agtr1 antagonist losartan decreased the phagocytic activity of Agtr1+/+ macrophages in vivo to a level comparable to that of Agtr1–/– macrophages.
Figure 8Phagocytic activity of Agtr1–/– and Agtr1+/+ macrophages. (a) Fluorescent beads were injected into the peritoneal cavity after macrophage induction by thioglycollate, and the exudate was analyzed by flow cytometry for in vivo assay of phagocytosis. Histograms include Agtr1+/+ macrophages, Agtr1+/+ macrophages treated with 10–6 M losartan, and Agtr1–/– macrophages. Three separate experiments were carried out, and representative results are shown. (b) Peritoneal macrophages were incubated in vitro with fluorescent beads, and the phagocytic activity was analyzed by flow cytometry. Histograms include fluorescent beads only, Agtr1+/+ macrophages incubated without fluorescent beads, and Agtr1+/+ and Agtr1–/– macrophages incubated with fluorescent beads. The mean fluorescence intensity level (lin Mean X) of each peak is shown for fluorescent beads only. The percentages of macrophages with multiple beads (more than eight) are shown next to the Agtr1+/+ and Agtr1–/– histograms, respectively. R1 denotes the peak derived from one fluorescent bead; R2, two beads; R3, three beads; and R4, four beads. Three separate experiments were carried out, and representative results are shown. (c) In vitro phagocytosis of fluorescent beads by Agtr1–/– (left) and Agtr1+/+ (right) macrophages viewed under a light microscope. Magnification, ×400. (d) In vivo phagocytosis of fluorescent beads by Agtr1–/– (left) and Agtr1+/+ (right) macrophages. Magnification, ×400.
In the in vitro phagocytosis assay, isolated macrophages were incubated with fluorescent beads, and the macrophages with fluorescent beads were counted by flow cytometry in accordance with the amount of ingested beads (33, 35). Representative results are shown in Figure 8b. There was no difference in the number of macrophages with beads (phagocytic macrophages) between Agtr1+/+ and Agtr1–/– macro-phages (84.9% ± 3.0% vs. 79.4% ± 2.6 % phagocytic macrophages, not significant; duplicate experiments, n = 6 per group). By contrast, there were significantly more beads in Agtr1+/+ macrophages than in Agtr1–/– macrophages (mean fluorescence intensity of total macrophages, 722.3 ± 53.9 vs. 498.4 ± 21.9, P < 0.005). There was no difference between Agtr1+/+ and Agtr1–/– macrophages in the number of macrophages with nonspecifically bound beads (8.9% of Agtr1+/+ macrophages and 10.4% of Agtr1–/– macrophages) or in the fluorescence intensity of nonspecifically bound beads (mean fluorescence intensity of total macrophages; 11.5 vs. 12.7, respectively). The Agtr1 antagonist losartan had no significant effect on either the number of the phagocytic macrophages (84.9% ± 3.0% vs. 79.5% ± 0.8%, not significant) or the number of phagocytosed beads (mean fluorescence intensity of total macrophages, 722.3 ± 53.9 vs. 642.0 ± 30.4, not significant). No significant difference was also observed between Agtr1+/+ macrophages with or without angiotensin II treatment at 10–6 M (86.4% ± 1.2% vs. 84.9% ± 3.0 % phagocytic macrophages, not significant; mean fluorescence intensity of total macrophages, 722.3 ± 53.9 vs.736.1 ± 35.8, not significant).
Together, these in vivo and in vitro analyses indicate that Agtr1+/+ macrophages are exposed to angiotensin II in vivo, exhibit elevated phagocytic capability, and, as a consequence, are not fully susceptible to either deactivation by losartan or further activation by angiotensin II in vitro.
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