Delay in hepatocyte proliferation and prostaglandin D2 synthase expression for cholestasis due to endotoxin during partial hepatectomy in rats
- Authors:
- Published online on: September 13, 2019 https://doi.org/10.3892/mmr.2019.10681
- Pages: 4367-4375
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Copyright: © Wakasa et al. This is an open access article distributed under the terms of Creative Commons Attribution License.
Abstract
Introduction
The liver is a unique organ with the capacity to regenerate following the removal of two-thirds of liver mass (1). Liver regeneration requires the precisely coordinated proliferation of the two major hepatic cell populations, hepatocytes and liver sinusoidal endothelial cells to reconstitute liver structure and function (2). Liver regeneration also requires the interaction between hepatocytes and other component cells, such as Kupffer cells and hepatic stellate cells (1,3,4). Numerous molecules, including hepatocyte growth factor and epidermal growth factor have been demonstrated as mitogens produced in nonparenchymal cells (5). Suppressed liver regeneration is of major concern for small remnant liver volume in adult living donor transplantation or in bacterial infection after partial hepatectomy (PH), as this has been associated with cholestasis and mortality (6).
Hepatocytes under physiological conditions efficiently extract bile acids from sinusoids via the sodium-dependent taurocholate cotransporting polypeptide (Ntcp) and the sodium-independent organic anion transporting polypeptide (Oatp1) (7). The extracted bile acids are excreted into the bile canaliculi by ATP-dependent transporters, such as the bile salt export pump (7). In our previous study, 90% PH in rats resulted in high blood bile acids levels and the suppression of Ntcp expression (6). Thus, lower uptake of bile acids has been suggested to be partly involved in cholestasis (6).
Infection is a frequent complication after living donor liver transplantation (8). Low-dose lipopolysaccharide (LPS) application after PH in mice was reported to delay liver proliferation (9). As LPS is known to activate Kupffer cells (10), this suggests that activated Kupffer cells may inhibit liver proliferation; however, it has been demonstrated that Kupffer cells stimulate liver regeneration after PH (1); depletion of Kupffer cells by clodronate delays liver regeneration (11). Therefore, Kupffer cells activated by LPS may lose their capacity to induce hepatocyte proliferation after PH.
The present study examined whether LPS-induced cholestasis is also due to the suppression of Ntcp expression, as observed in 90% PH rats. It also examined whether Kupffer cells activated by LPS inhibit or stimulate liver regeneration after PH. The expression of anion transporters for the uptake from the sinusoid was decreased in PH, but LPS did not further decrease their expression. This suggested that decreases in these transporters were not responsible, but a delay in hepatocyte proliferation may be linked to LPS-induced cholestasis. LPS treatment alone or in combination with PH induced Kupffer cell activation with a CD163-positive phenotype, a marker for M2-type macrophages (12); CD163-positive cells were suggested to produce chemokine ligand 9 (Cxcl9), which was determined to be involved in chronic inflammation (13) and M2 macrophage polarization (14). As hematopoietic type prostaglandin D2 synthetase (Ptgds2) is known to inhibit lymphocyte proliferation (15), Ptgds2 staining was performed. Hepatocytes in the LPS + PH group were stained and markedly stained at 24 h, a time point when cell proliferation was notably inhibited. On the contrary, hepatocytes in the LPS or the PH groups were not stained.
Materials and methods
Animals and animal treatment
Male Sprague-Dawley rats weighing 180–220 g and 6 weeks old were purchased from Charles River Laboratories Japan, Inc. In total 39 rats were used and they were housed under routine laboratory conditions at the animal laboratory of Hirosaki University. The rats received standard laboratory chow, had free access to food and water, and were kept in a thermostatically controlled room (25°C) with a 12-h light-dark cycle. Before undergoing surgical procedures, all rats were fasted for 24 h. The rats were divided into five groups: Control group without any treatment, sham group receiving laparotomy alone, LPS group receiving intravenous LPS 75 µg/rat, PH group receiving 70% PH, and LPS + PH group receiving intravenous LPS injection immediately after PH. 70% PH was performed as reported previously (6). The rats of four groups except the control group were sacrificed at 24, 72 and 168 h after laparotomy or PH and/or LPS treatment. Those of the control group were sacrificed at 0 h. Three rats each were used at respective time points of each group. LPS (O55:B5, L2880) was purchased from Sigma-Aldrich (Merck KGaA). After the surgical procedures, the rats had free access to a 200 g/l glucose solution for 24 h to avoid post-operative hypoglycemia after hepatectomy. The present study was performed in accordance with the Guidelines for Animal Experimentation, Hirosaki University, and all of the animals received humane care according to the criteria outlined in the ‘Guide For The Care And Use Of Laboratory Animals’ prepared by the National Academy of Sciences and published by the National Institutes of Health (16).
Plasma total bilirubin and bile acids
Blood from the hearts was collected in test tubes containing EDTA and plasma was prepared after centrifugation at 2,500 × g, for 10 min at room temperature. Plasma total bilirubin, aspartate aminotransferase (AST), and alanine aminotransferase (ALT) were measured using Spotchem EZ (ARKRAY, Inc.) with SPOTCHEM II Basic Panel 2 Test Strips (MT-7785; ARKRAY, Inc.), according to the manufacturer's protocols. The plasma levels of total bile acids were measured with an assay kit (Diazyme Laboratories), according to the manufacturer's protocol.
Microarray analysis
Total RNA was extracted from frozen liver samples at 0, 24, 72 and 168 h after 70% hepatectomy and/or LPS injection with TRIzol® reagent (Thermo Fisher Scientific, Inc.). Equal amounts of RNA from three individual livers were combined, and 10 µg of RNA was used to produce biotin-labeled complementary RNA (cRNA) with GeneChip IVT labeling kit (Affymetrix; Thermo Fisher Scientific, Inc.). The labeled and fragmented cRNA was subsequently hybridized to GeneChip® Rat Gene-ST 2.0 Array (Affymetrix; Thermo Fisher Scientific, Inc.). Labeling, hybridization, image scanning and data analysis were performed at TOHOKU CHEMICAL Co., Ltd.
Reverse transcription-quantitative polymerase chain reaction (RT-qPCR)
Complementary DNA (cDNA) was reverse-transcribed from 1 µg of total RNA using the Omniscript RT kit (Qiagen, Inc.), according to the manufacturer's protocols. A MiniOpticon Detection System (Bio-Rad Laboratories, Inc.) and SYBR® Green Supermix (Bio-Rad Laboratories, Inc.) were used for the quantitation of specific mRNA. The amplification of ubiquitin C cDNA was performed to standardize the levels of the target cDNA, as reported previously (6). Gene-specific primers were designed according to known rat sequences (Table I). PCR amplification consisted of 30 sec at 94°C, 30 sec at 55–60°C and 30 sec at 72°C for 30–35 cycles. No non-specific PCR products, as detected by melting temperature curves, were found. After normalizing the expression of the target gene to ubiquitin C expression using the 2−ΔΔCt method reported by Livak and Schmittgen (17) in triplicate; the levels of mRNA expression in three samples at respective time points (0, 24, 72, and 168 h after treatment) were expressed relative to the control values.
Western blotting
Crude liver membranes were prepared according to the method of Gant et al (18) and the samples (100 µg protein each) were dissolved in sample buffer and separated via 7.5% SDS-PAGE with a 4.4% stacking gel. Protein content was measured by Bradford's method (19) using a bovine serum albumin standard curve. Following electrophoresis, the proteins were transferred to polyvinylidene fluoride membranes (Hybond-P, GE Healthcare). After blocking with 4% nonfat dry milk in Tris-buffered saline for 2 h at room temperature, membranes were incubated overnight at 4°C with primary anti-Ntcp antibody (sc-107029; 1:10,000, Santa Cruz Biotechnology, Inc.) or anti-β-actin antibody (ab227387; 1:1,000, Abcam). Immune complexes were detected using a horseradish peroxidase conjugated anti-rabbit IgG secondary antibody (NA934; 1:2,000, GE Healthcare) and visualized with an enhanced chemiluminescent kit (ECL Plus; GE Healthcare).
Immunostaining
Liver tissue samples were fixed in 10% neutral buffered formaldehyde for two days at 4°C and embedded in paraffin. These paraffin blocks were sliced into 4 µm sections and passed through xylene and a graded alcohol series. The deparaffinized sections were stained with hematoxylin solution at room temperature for 5 min. Following washing with water and passing through a graded alcohol series, the sections were stained with eosin solution for 1 min. The deparaffinized sections were also stained for CD68, CD163, Cxcl9, and Ptgds2 using a standard avidin-biotin-peroxidase conjugate method (20) using an automated immunostaining instrument (Benchmark XT; Ventana Medical System). The slides were blocked with 0.3% hydrogen peroxide and then incubated for 1 h at room temperature with the primary antibodies. The antibodies employed were: Anti-CD68 antibody (MCA 341R; 1:100, Bio-Rad Laboratories, Inc.), anti-CD 163 antibody (sc-58965; 1:500, Santa Cruz Biotechnology, Inc.), anti-Cxcl9 antibody (bs-2551R; 1:500, BIOSS Inc.), and anti-Ptgds2 antibody (PA 5-43217; 1:500, Invitrogen; Thermo Fisher Scientific, Inc.). Non-immune γ-globulin fractionated from rabbit sera by 20–40% saturation of ammonium sulfate (21) was used as a negative control instead of primary antibody. The biotinylated anti-rabbit IgG or anti-mouse IgG antibodies and Vectastain ABC kit (PK6101) were obtained from Vector Laboratories, Inc. The specific binding was visualized with a 3,3′-diaminobenzidine tetrahydrochloride solution. Sections were then lightly counterstained with hematoxylin for microscopic examination. Images were captured with an inverted FSX 100 microscope (Olympus Corporation). Digital images were processed with Adobe Photoshop (version 7.0, Adobe Systems, Inc.) and ImageJ software (v1.50, National Institutes of Health).
Statistical analysis
Experiments for which a statistical analysis was indicated were performed independently at least three times. Data are presented as the mean ± standard deviation. Statistical comparisons were analyzed using SPSS software (v22.0, IBM Corp.). Differences between experimental groups were assessed for significance using two-way ANOVA with a Tukey's post-hoc test. P<0.05 was considered to indicate a statistically significant difference.
Results
Elevated plasma bilirubin and bile acid levels in the LPS + PH group
Bilirubin and bile acid levels in the plasma at 24 h post-operation were significantly increased in the LPS + PH group compared with those in the sham group. The bile acid level was significantly higher in the LPS + PH group than that in the PH group (Fig. 1). These results indicated that LPS induced cholestasis in this rat model. AST and ALT levels in the plasma at 24 h were significantly increased in the LPS + PH group and PH group, compared with those in the sham group.
Suppression and delay in DNA replication in the LPS + PH group
Microarray analysis was performed to comprehensively analyze alterations in liver gene expression. Data were expressed as signal values, and changes of >2-fold or <1/2 from the values in the control or sham groups were considered significant. Ribonucleotide reductase regulatory subunit M2 (Rrm2), DNA topoisomerase IIα and proliferating cell nuclear antigen (Pcna), which are markers of DNA replication (6), reached a peak level of expression after 24 h in the PH group and gradually decreased thereafter. However, in the LPS + PH group, these replication signals were low after 24 h and peaked after 72 h. The values at 72 h were lower than those at 24 h in the PH group (Table II). These results suggested a delay and suppression in DNA replication in the LPS + PH group. No notable changes were observed in Cd68 or Cd163 expression, which are markers of Kupffer cells (12,22). The chemokine Cxcl9 markedly increased in the LPS group and LPS +PH group, compared with that in the sham at 24 h (Table II). For sinusoid transporters, Ntcp (Slc10a1), Oatp1(Slc21a1) and Oatp2 (Slc21a2) were reduced in the LPS + PH and the PH groups at 24 h. These expression levels returned to control levels at 72 h in both groups. No notable changes were observed in collagen 1α1 or desmin, markers of hepatic stellate cells, or in cytokeratin19 or epithelial cell adhesion molecule, markers of liver progenitor cells (23) (Table II).
To confirm these changes in gene expression, RT-qPCR was performed. Abcc2, Oatp1, and Oatp2 mRNA levels were significantly decreased at 24 h in the LPS + PH group and PH group, compared with those in the sham group. These mRNA levels except Oatp2 were not significantly different between the LPS + PH and the PH groups (Fig. 2A). The Rrm2 mRNA levels at 24 h in the LPS + PH group were lower than those in the PH group (Fig. 2B). Rrm2 and Pcna peaked at 24 h in the PH group, whereas at 72 h, the levels increased in the LPS + PH group (Fig. 2B), confirming the results obtained by microarray analysis. Cxcl9 showed a significant rise after 24 h in the LPS and LPS + PH groups compared with the control and PH groups, respectively (Fig. 2B). These findings suggested that Cxcl9 expression was dependent on LPS treatment.
Ntcp protein levels were examined by western blotting; Ntcp expression was decreased in the PH and LPS + PH group at 24 h compared with the sham group (Fig. 3).
Expression of Cxcl9 in Kupffer cells activated by LPS treatment
Although Cd68 mRNA or Cd163 mRNA levels were unaltered as determined by microarray analysis (Table II), staining for CD68, a marker of Kupffer cells and macrophages (22), revealed a marked increase in CD68-positive Kupffer cells in the LPS and LPS + PH groups, compared with that in the sham and PH groups (Fig. 4A). CD163 staining, a marker for M2 macrophages and Kupffer cells (12) was positive in cells in the LPS and LPS + PH groups (Fig. 4B). These CD163-positive cells were not detected in the sham or PH groups. There were fewer CD163-positive cells than CD68-positive cells, and their cell shapes were different from each other. These results suggested that CD163-positive cells detected after LPS treatment denoted M2-type Kupffer cells (12). There were also Cxcl9-positive cells in the LPS and LPS + PH groups (Fig. 4C), whereas Cxcl9-positive cells were not detected in the sham or PH groups. The number of Cxcl9-positive cells was similar to that of CD163-positive cells rather than CD68-positive cells (Fig. 4D).
Expression of Ptgds2 in hepatocytes in the LPS + PH group
As Ptgds2 inhibits cell proliferation (15), Ptgds2 staining was performed. A positive reaction was only detected in hepatocytes of the LPS + PH group, but not in other groups (Fig. 5). Kupffer cells were not stained in any groups. In the LPS + PH group, Ptgds2 was markedly stained in hepatocytes at 24 h, weakly stained at 72 h, but not at 168 h.
Discussion
In the rat PH model of the present study, LPS treatment induced cholestasis and delayed cell proliferation. Compared with the sham group, the expression of anion transporters involved in the uptake from the sinusoid was downregulated at 24 h in both the PH and the LPS + PH groups, but did not differ between the latter two groups. Downregulation of these anion transporters is a causative factor for cholestasis after 90% PH (6,7,24). However, this was unlikely in the case of cholestasis in the LPS + PH group; suppression or delays in cell proliferation may be the responsible factor. Downregulation of marker genes of DNA replication, such as Rrm2 was determined by RT-qPCR analysis; however, delays in cell proliferation are not confirmed by protein levels, as immunohistochemistry for Pcna was not conducted. Hepatocyte proliferation is blocked by 2-acetylaminofluorene administration during PH in rats (25). In this case, biliary epithelial cells and hepatic stellate cells become progenitor cells, and these cells contribute to liver regeneration (25). In the case of LPS, activation of these cells was not detected, and microarray and RT-qPCR data suggested that hepatocyte proliferation was inhibited transiently.
Our findings revealed that LPS treatment increased the count of CD68-positive cells and CD163-positive cells. These results confirmed the activation of Kupffer cells by LPS as reported previously (26). From microarray analysis, Cd68 and Cd163 expression was determined to be unaffected by LPS treatment despite increases in the number of CD68- and CD163-positive cells as detected by immunostaining. This discrepancy may reflect a difference between mRNA and protein expression; however, further investigation is required. As CD163 is a marker of M2-type macrophages (12,22), CD163-positive cells may belong to M2-Kupffer cells (22). Thus, CD68-positive cells may denote M1-type macrophages or Kupffer cells (27). A marked increase in the number of CD68-positive cells by LPS treatment raises two possibilities: The proliferation of CD68-positive cells in the liver or the migration of CD68-positive cells to the liver from bone marrow (28,29). The absence of alterations in Cd68 mRNA levels by LPS treatment suggests the latter explanation as a more likely possibility.
In the present study, Cxcl9 was significantly induced by LPS treatment. Immunohistochemistry suggested that Cxcl9 was expressed by CD163-positive cells. Double staining for Cxcl9 and CD163 should be conducted to establish this possibility. Cxcl9 is a member of a family of ligands for the Cxcr3 receptor, which is involved in chronic inflammation and cancer (13). Cxcl9 is also a biomarker of acute cellular rejection after liver transplantation (30). Endothelial cell growth is stimulated or inhibited depending on alternatively spliced variants of Cxcr3 (31). Cxcl9 is expressed in macrophages (32,33) and C-X-C motif chemokine receptor 3 (Cxcr3) promotes M2 macrophage polarization in human liver cancer (14). Prostaglandin E2 inhibits CXCR3 ligand secretion induced by interferon-γ treatment in human breast cancer cells (34).
Ptgds2 is the hematopoietic-type Ptgds and is expressed in mast cells and macrophages (35). Ptgds2 is also expressed in skeletal muscle cells with muscular dystrophy (36). Inhibition of Ptgds2 stimulates the survival of muscle cells via the suppression of muscular cell death (37). Lymphocytes isolated from Ptgds2 knock-out mice exhibit hyperproliferation (15). The time courses of Ptgds2 staining and cell proliferation had opposite profiles in our study. Thus, Ptgds2 was suggested to suppress hepatocyte proliferation. Ptgds2 was not detected in the LPS or PH groups, but was expressed in hepatocytes of the LPS + PH group. These results indicated that LPS and cell proliferation signals may be required for the induction of Ptgds2 expression in hepatocytes. The findings indicating that LPS alone did not alter cell proliferation suggested that a delay in cell proliferation in the LPS + PH group may not be due to the direct effects of LPS on hepatocytes, but due to Kupffer cells activated by LPS. Cxcl9 may be a candidate signaling molecule released from Kupffer cells for Ptgds2 expression in hepatocytes; however, because Cxcl9 was produced by LPS alone, Cxcl9 may not be sufficient for Ptgds2 expression. Ptgds2 may be a target to prevent a delay in cell proliferation after PH induced by LPS or bacterial infections.
Acknowledgements
The authors would like to thank Ms. Yukie Fujita and Ms. Sayumi Kubo (Department of Pathology and Bioscience, Hirosaki University Graduate School of Medicine) for their technical assistance in immunostaining, and Ms. Ryoko Seito, Mr. Hitoshi Kudo and Ms. Ikumi Shirahama (Institute for Animal Experimentation, Hirosaki University Graduate School of Medicine) for their technical assistance for animal treatment.
Funding
The present study was partly supported by the Japan Society for the Promotion of Science KAKENHI (grant no. 15K20845; Grant-in-Aid for Young Scientists B). No additional external funding was received for this study.
Availability of data and materials
All data generated or analyzed during the present study are included in this published article.
Authors' contributions
YW, NK and ST conceived the idea and design of the present study. YW, NK, TY and TS performed the animal experiments. YW, KH and ST wrote the manuscript. YW, NK and ST discussed the results and contributed to the final version of the manuscript. KH performed the statistical analyses. All authors approved the final version of the manuscript to be published.
Ethics approval and consent to participate
All animal experiments were conducted strictly according to ethical standards and approved by the Animal Ethical Committee of Hirosaki University Graduate School of Medicine (approval ID: M15041).
Patient consent for publication
Not applicable.
Competing interests
The authors declare that they have no competing interests.
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