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. Author manuscript; available in PMC: 2021 Jul 16.
Published in final edited form as: Toxicol Lett. 2011 Jan 13;201(3):213–220. doi: 10.1016/j.toxlet.2011.01.002

Lipopolysaccharide down-regulates carbolesterase 1 and 2 and reduces hydrolysis activity in vitro and in vivo via p38MAPK -NF-κB pathway

Zhao Mao 1,2,#, Yang Li 1,#, Yan Peng 1, Xiaofei Luan 1, Haiyan Gui 1, Xuemin Feng 1, Gang Hu 1, Jianping Shen 1, Bingfang Yan 3, Jian Yang 1,*
PMCID: PMC8284319  NIHMSID: NIHMS277491  PMID: 21237253

Abstract

Carboxylesterases constitute a class of enzymes that hydrolyze drugs containing such functional groups as carboxylic acid ester, amide, and thioester. Hydrolysis of many drugs is reduced in liver diseases such as hepatitis and cirrhosis. In this study, we have demonstrated, in vitro and in vivo, treatment with LPS decreased the expression of HCE1 and HCE2 and the capacity of hydrolytic activity. In HepG2 cells, the decreased expression by LPS occurred at both mRNA and protein levels. Both HCE1 and HCE2 promoters were significantly repressed by LPS, and the repression was comparable with the decrease in HCE1 and HCE2 mRNA, suggesting the transrepression is responsible for suppressed expression. Further study showed that both PDTC, a NF-κB inhibitor, and SB203580, a p38MAPK inhibitor, could abolish the repression of HCE1 and HCE2 mediated by LPS, but U0126, a selective ERK1/2 inhibitor, could not do so, suggesting the repression of HCE1 and HCE2 by LPS through the p38MAPK-NF-κB pathway. In addition, being pretreated with LPS, HepG2 cells altered the cellular responsiveness to ester therapeutic agents, including clopidogrel (hydrolyzed by HCE1) and irinotecan (hydrolyzed by HCE2). The altered cellular responsiveness occurred at low micromolar concentrations, suggesting that suppressed expression of carboxylesterases by LPS has profound pharmacological and toxicological consequences, particularly with those that are hydrolyzed in an isoform-specific manner. This study provides new insight into the understanding of the pharmacological and toxicological effects and the mechanisms for repressing drug metabolism enzymes in inflammation.

Keywords: lipopolysaccharide (LPS), carboxylesterases1 (HCE1), carboxylesterases2 (HCE2), p38MAPK, NF-κB

Introduction

The liver is the richest source of drug-metabolizing enzymes in terms of abundance and diversity, thereby playing a determinant role in drug metabolism (Parkinson, 2001). Although many factors may alter the hepatic capacity of drug, regulated expression of drug-metabolism enzymes contributes the most to the alteration (Parkinson, 2001; Poso and Honkakoski, 2006). Transactivation by nuclear receptors such as the pregnane X receptor is largely responsible for the increased expression of these genes (Poso and Honkakoski, 2006). However, the mechanisms of the down-regulation remain largely unclear.

Carboxylesterases constitute a class of enzymes that hydrolyze drugs containing such functional groups as carboxylic acid ester, amide, and thioester (Satoh and Hosokawa, 2006). The liver expresses two major carboxylesterases, including human carboxylesterases1 (HCE1) and carboxylesterases2 (HCE2), whereas the gastrointestinal tract expresses predominantly HCE2 (Schweretal., 1997; Satoh and Hosokawa, 2006). In addition to the difference in tissue distribution, these two enzymes differ markedly in the hydrolysis of certain drugs. For example, HCE1 but not HCE2 rapidly hydrolyzes antithrombogenic agent clopidogrel (Tang et al., 2006), while, HCE2 but not HCE1 rapidly hydrolyzes anticancer agent irinotecan (Wu et al., 2002; Yang et al., 2007).

Hydrolysis of many drugs is reduced in liver diseases such as hepatitis and cirrhosis (Thiollet et al., 1992; Gross et al., 1993). For example, the hydrolysis of perindopril, a nonsulfhydryl angiotensin-converting enzyme inhibitor, is decreased by as much as 50% in patients with cirrhosis (Thiolletetal., 1992). Likewise, the hydrolysis of cilazapril, another anti-hypertensive, is decreased by 45% in hepatitis patients (Grossetal.,1993). The bacterial endotoxin, lipopolysaccharide (LPS), induces the expression of proinflammatory cytokines such as tumor necrosis factor-α (TNFα) and interleukin-6 (IL-6) in the liver, which contribute to altered DMEs expression. (Myers et al., 2010). It can reduce the CYP3A4 in human hepatocytes (Gu et al., 2006) or CYP3A11 in mice (Ghose et al., 2004).

In this study, we have demonstrated in HepG2 cells that treatment with LPS decreased the expression of HCE1 and HCE2. The decreased expression occurred at both mRNA and protein levels, and it was confirmed by enzymatic assay. Both HCE1 and HCE2 promoters were significantly repressed by LPS, and the repression was comparable with the decrease in HCE1 and HCE2 mRNA. In addition, being pretreated with LPS, HepG2 cells altered the cellular responsiveness to ester therapeutic agents, including clopidogrel and irinotecan. The altered cellular responsiveness occurred at low micromolar concentrations, suggesting that suppressed expression of carboxylesterases by LPS has profound pharmacological and toxicological consequences.

Materials and methods

Chemicals and supplies

Lipopolysaccharide (LPS) (Escherichia coli 055:B5), ammonium pyrrolidinedithiocarbamate (PDTC), 4-(4-Fluorophenyl)-2-(4-methylsulfinylphenyl)-5-(4-pyridyl)-1H-imidazole (SB203580), and 1,4-Diamino-2,3-dicyano-1,4-bis(o-aminophenylmercapto)butadiene monoethanolate (U0126) were purchased from Sigma (St. Louis, MO, USA). Dulbecco's modified Eagle's medium (DMEM), Trizol and high-fidelity platinum Taq DNA polymerase were from Invitrogen (Carlsbad, CA, USA). Trypase, MLV reverse transcriptase, RNase inhibitor, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium (MTT), and Dual-luciferase reporter assay system were from Promega (Madison, WI, USA). GeneJet™ DNA VitroTransfection Reagent(Ver II) was from SignaGen Labortories (Gaithersburg, MD, USA). Fetal bovine serum was from Sijiqing (Hangzhou, China). The antibody against glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and anti- carboxylesterase 1 (CSE1) were from Abcam (Cambridge, UK). The anti- carboxylesterase 2 (CSE2) was from Abgent (San Diego, CA, USA). The goat anti-rabbit IgG conjugated with horseradish peroxidase was from Pierce Chemical (Pierce, Rockford, IL, USA). Nitrocellulose membrane was from Bio-Rad Laboratories (Hercules, CA, USA). All other reagents were from sigma (St. Louis, MO, USA).

The culture and treatment of HepG2 cells

Hepatoma (HepG2) cells were purchased from American Type Culture Collection (Mannassas, VA, USA), and maintained in the Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum, penicillin/streptomycin, and 1×nonessential amino acids. HepG2 cells were seeded at the density of 5×106 cells/well (6-well plates for protein level), of 5×105 cells/well (12-well plates for mRNA level) or of 8×104 cells/well (48-well plates for reporter assay) in a regular medium. The cells were treated with 5 μg/ml LPS for 0, 3, 6, 9, 12, 24 h (for mRNA level) or 0.1, 1, 5 μg/ml LPS for 48 h (for protein level and enzymatic assay) (Yang and Yan, 2007), and the treated cells were cultured in a 1% serum-reduced medium.

Quantitative reverse transcription-polymerase chain reaction

Total RNA was isolated by using a Trizol according to the manufacturer's instruction and checked by formaldehyde gel electrophoresis for quality control. The first-strand cDNA was synthesized using total RNA (1 μg) at 25〈 for 10 min, 42〈 for 50 min, and 70〈 for 10 min by using random primers and moloney murine leukemia virus reverse transcriptase (Promega, Madison, WI,USA). The cDNAs were then diluted eight times and quantitative PCR was conducted with TaqMan Gene Expression Assay kits (Applied Biosystems, Foster City, CA, USA). The TaqMan assay identification numbers are: HCE1, Hs00275607_m1; HCE2, Hs00187279_m1; GAPDH, 4352934E; mCE1, Mm00491334_m1; mCE2, Mm00524035_m1; mGAPDH; Mm99999915_g1. 20 μl PCR mix contained 10μl of universal PCR master mixture, 1 μl of gene-specific TaqMan assay mixture (probe), 6 μl of diluted cDNA as template and 3 μl of water. The PCR amplification and quantification were done in an Applied Biosystems 7900 real-time PCR system (Applied Biosystems, Foster City, CA, USA) with one cycle at 50〈 for 2 min and 95〈 for 10 min, followed by 40 cycles of 95〈 for 15 s and 60〈 for 1 min. The signals from each target gene were normalized based on the signal from corresponding GAPDH.

Reporter constructs

The HCE2 promoter reporter (−1931/ +6) was kindly provided by Dr. M.E. Dolan (University of Chicago, Chicago,IL) (Wu et al.,2003). HCE1A2 promoter reporter was prepared as previously described (Yang et al., 2007). The sequences of constructs were verified by direct DNA sequencing.

Transient co-transfection experiment

HepG2 cells were plated in 48-well plates in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum at the density of 8×104 cells/well. Transfection was conducted by GeneJet™ (Ver II)(Gaithersburg, MD, USA). Its mixtures contained 50 ng of a reporter plasmid, and 5 ng of Null-Renilla reniformis luciferase plasmid. HepG2 cells were transfected for 12 h and the medium was replaced with fresh medium supplemented with 1% fetal bovine serum with or without LPS (5 μg/ml). After another 24 h of treatment, the cells were washed once with phosphate-buffer saline (PBS) and collected by scraping. The collected cells were shaken for 20 min in room temperature. The reporter enzyme activities were assayed with Dual-Luciferase reporter assay system which used two substrates, the firefly luminescence and Renilla luminescence to determine the activities of two luciferases sequentially. The firefly luciferase activity, which reflected the reporter activity, was evaluated by mixing an aliquot of lysates (10 μl) with Luciferase Assay Reagent II (Promega, Madison, WI). Then, the firefly luminescence was quenched and the Renilla luminescence was activated simultaneously by adding Stop & Glo reagent (Promega) to the sample tubes. The firefly luminescence signal intensity was normalized based on the intensity of Renilla luminescence signal, and the ratio of normalized luciferase activity from LPS over PBS treatment served as the relative luciferase activity.

Mice and treatment

Male CD1 mice (16-week old) were obtained from the experimental animal center of Nanjing (Nanjing, China). Mice were kept under environmentally controlled conditions (ambient temperature, 22°C; humidity, 40%) on a 12 h light/dark cycle with food and water adlibitum. Male mice were injected intraperitoneally with LPS (5 mg/kg; Sigma Chemicals, St. Louis, MO, USA) in saline (Sachdevak et al., 2003), and the controlled mice were treated with saline only. At 24 h after injection for sacrifice, mice were i.p. injected with ketamine (1 ml/kg at 100 mg/ml). When mice were completely anesthetized (~5 min), surgery was performed to expose the liver. The liver was perfused with PBS through the portal vein to remove blood. The upper intestine of mice was isolated and washed with cold PBS. The perfused liver and intestine were then divided into two parts, with one part being immediately used for preparing total RNA and the other one frozen at −80°C for preparing S9 fractions. The use of animals was approved by IACUC (Institutional Animal Care and Use Committee) of Nanjing Medical University. And every effort was made to minimize animal suffering and to reduce the number of animals used for experiments.

Preparation of S9 fractions

The frozen livers and intestines were thawed in homogenization buffer (50 mM Tris–HCl, pH 7.4, 150 mM KCl and 2 mM EDTA) and then homogenized with 6 passes of Teflon pestle driven by a Wharton stirrer. The homogenates were centrifuged at 10,000×g for 20 min at 4°C. The S9 fractions of livers and intestines (supernatant) were assayed for the hydrolysis of para-nitrophenylacetate and for Western Blot.

Western analysis

HepG2 lysates (20 μg) or S9 fractions of liver and intestine (10 μg) were resolved by 7.5% SDS- polyacrylamide gel electrophoresis and electrophoretically transferred to a nitrocellulose membrane. After nonspecific binding sites were blocked with 5% nonfat milk, the blots are incubated with an antibody against HCE1 (1:2500), HCE2 (1:2500) and GAPDH (1:4000). The primary antibodies were subsequently localized with goat anti-rabbit IgG conjugated with horseradish peroxidase. Horseradish peroxidase activity was detected with a chemiluminescent kit (Pierce, Rockford, IL, USA). The chemiluminescent signal was captured by KODAK Image Station 2000 (Estman Kodak, Rochester, NY, USA), and the bands density was quantified by KODAK Image Analysis software (Estman Kodak, Rochester, NY, USA), and expressed as HCE1 or HCE2/GAPDH arbitrary units.

Enzymatic assay

HepG2 cells were treated with LPS as described above. Cells were rinsed with PBS and harvested in 300 μl of 100 mM potassium phosphate buffer, pH 7.4. The cell suspension was sonicated by a sonifier (Nanjing, China), and the cell debris was removed by centrifugation at 12,000 g for 15 min at 4〈. The supernatant or S9 fractions of liver and intestine were assayed for hydrolytic activity toward para-nitrophenylacetate as described previously (Yang and Yan, 2007). A sample cuvette (1 ml) contained 10 μg of cell lysates or S9 fractions in 100 mM potassium phosphate buffer pH 7.4, and 1 mM substrate at room temperature. Reactions were initiated by adding para-nitrophenylacetate (10 μl of 100 mM stock in acetonitrile), and the hydrolytic rate was recorded from an increase in absorbance at 400 nm. The extinction coefficient (E400) was determined to be 13 mM−1 cm−1. Several controls were performed, including incubation without proteins.

Cytotoxicity assay

HepG2 cells were seeded into 96-well plates at the density of 5000 cells/well. After an overnight incubation, the medium was replaced with fresh medium with or without 5 μg/ml LPS in1% FBS DMEM. After an additional 12-h incubation, the cells were washed with DMEM once and treated with clopidogrel or irinotecan at various concentrations. When the cells were treated for 48 h, the medium was replaced with fresh medium containing 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium(MTT) at a final concentration of 0.5 mg/ml. After 2-h incubation at 37〈, the medium was gently decanted, and dimethysulfoxide (100 μl/well) was added to dissolve formazan product. The optical density (OD) was determined at 570 nm, and the final OD values were expressed by subtracting the background reading (no seeded cells). And morphologic changes were detected under microscope before MTT assay.

Other analyses

Protein concentrations were determined with BCA protein assay (Pierce Chemical) based on albumin standard. Data are presented as mean ± S.D. of at least three separate experiments, except where results of blots are shown, in which case a representative experiment is depicted in the figures. Statistical analysis was performed using SPSS 10.0 for Windows (SPSS Inc, Chicago, IL, USA). And statistical analysis for multiple comparisons was performed by a one-way ANOVA test with Bonferroni's corrections. The differences were considered statistically significant when P < 0.05.

Results

LPS reduces the overall hydrolytic activity by decreasing the HCE1 and HCE1 expression in HepG2 cells

It has been reported that hydrolytic biotransformation decreased in patients with liver diseases such as hepatitis and cirrhosis (Thiollet et al., 1992; Gross et al., 1993; Prandota, 2005). To link the inflammation to the reduced hydrolysis, we tested whether LPS suppresses the expression of human carboxylesterases 1 and 2 (HCE1 and HCE2), the two major hydrolytic enzymes in the liver. HepG2 cells were cultured and treated with LPS for 24 h (mRNA level) or 48 h (protein level). Total RNA was prepared, and the levels of HCE1 and HCE2 were determined by qRT-PCR with TaqMan probes. HCE1 has two closely related genes that encode almost identical transcripts, and the HCE1 TaqMan probe recognized both transcripts. As shown in Fig.1A, the cell treated with LPS consistently decreased the HCE1, HCE2 mRNA level in a time-dependent manner. Next, we examined whether the decreases of HCE1 and HCE2 mRNA translate into the decreases in the hydrolytic activity. HepG2 cells were treated with LPS, and cell lysates were prepared. The overall hydrolytic activity by the lysates was determined with standard substrate para-nitrophenylacetate. Both HCE1 and HCE2 have been shown to hydrolyze this ester (Xie et al., 2002). Consistent with the decreases in HCE1 and HCE2 mRNA, the hydrolysis of para-nitrophenylacetate significantly decreased (Fig.2B). To determine whether the decreased hydrolysis is due to the decreased enzyme proteins, the lysates (20 μg) were analyzed by Western blotting for the abundance of HCE1 and HCE2. As expected, the levels of both HCE1 and HCE2 proteins significantly decreased in a dose-dependent manner, and the decrease was comparable with the that in the hydrolytic activity (Fig.1B, 1C).

Fig. 1. Effect of LPS on the levels of HCE1 and HCE2 in HepG2 cells.

Fig. 1

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(A) Effect of LPS on the levels of HCE1 and HCE2 mRNA in a time-dependent manner. HepG2 cells were seeded at the density of 5×105 in 12 well-plate overnight and the cells were treated with LPS (5 μg/ml) or the same volume of PBS for 0, 3, 6, 9, 12, 24 h. Total RNA was isolated and subjected to qRT-PCR analysis for the levels of HCE1 and HCE2 as described under Materials and Methods. The qPCR Cts were 23 for HCE1, 25 for HCE2 and 20 for GAPDH. All experiments were repeated at least three times, and the data were expressed as mean ± SD. * p<0.05, vs 0 h. (B) Effect of LPS on the hydrolytic of para-nitrophenylacetate. (C) Effect of LPS on the levels of HCE1 and HCE2 protein in a dose-dependent manner. HepG2 cells were seeded at the density of 5×106 in 6 well-plate overnight and the cells were treated with LPS (0.1, 1, 5 μg/ml) or the same volume of PBS for 48 h. Cell lysates were prepared and lysates (20 μg) were subjected to Western analyses with an antibody against HCE1, HCE2 or GAPDH. The relative protein level was expressed as HCE1 or HCE2/GAPDH arbitrary units, and the control (PBS) is considered as 1. Cell lysates (treated with 5 μg/ml) were assayed for the hydrolytic activity as described in Materials and Methods. All the experiments were repeated at least three times, and the data were expressed as mean ± SD. * p<0.05, a statistically significant decrease by LPS treatment.

Fig. 2. Effect of LPS on the cellular responsiveness to clopidogrel (A), and irinotecan (B).

Fig. 2

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HepG2 cells were seeded into 96-well plates at the density of 5000 cells/well. After an overnight incubation, 5 μg/ml LPS in 1% serum medium was added to half of the wells, and the treatment lasted for 12 h. Thereafter, the cells were washed with DMEM, and then they were treated with clopidogrel, or irinotecan at various concentrations. After an additional 48-h incubation (the medium was replaced with fresh drug-containing medium at 24 h), MTT was added to each well at a final concentration of 0.5 mg/ml. After 2-h incubation at 37°C, the medium was gently decanted, and dimethysulfoxide (100 μl/well) was added to dissolve formazan product. The OD was determined at 570 nm, and the final OD values were expressed by subtracting the background reading (no seeded cells). All the experiments were repeated at least three times, and data were expressed as mean ± SD. * p< 0.05, vs control group (clopidogrel, or irinotecan concentration is 0); # p<0.05, vs non-pretreatment group.

Alteration of cellular responsiveness to clopidogrel and irinotecan after pre-treatment with LPS

We next examined the pharmacological and toxicological consequences of the reduced expression of carboxylesterases by LPS. It has been reported that HCE1 and HCE2 differ markedly in the hydrolysis of antiplatelet agent clopidogrel and anticancer agent irinotecan (Tang et al., 2006; Wu et al., 2002). HCE1 rapidly hydrolyzes clopidogrel, whereas HCE2 rapidly hydrolyzes irinotecan. More importantly, hydrolysis of clopidogrel represents detoxication, whereas hydrolysis of irinotecan enhances cytotoxicity(Tang et al., 2006; Wu et al., 2002). It was assumed that LPS pre-treatment would increase cytotoxicity of clopidogrel, and the opposite would be true with irinotecan. HepG2 cells were first treated with 5 μg/ml LPS for 12 h, washed once, and then treated with clopidogrel or irinotecan at various concentrations. After an additional 48-h incubation, the cell viability was determined by MTT assay. And morphologic changes were detected under microscope before MTT assay. As shown in Fig.2, the cells pretreated with LPS alone caused no difference in the cell viability (The concentration of either clopidogrel or irinotean is 0) compared with those not pretreated with LPS. However, cells pretreated with LPS followed with clopidogrel or irinotecan represented statistically significant changes at corresponding concentration compared with those not pretreated with LPS (Fig.2). For example, the cells pretreated with LPS exhibited a significant decrease in cell viability when exposed to clopidogrel at as low as 1 μM, while the cells not pretreated with LPS exhibited a significant decrease in cell viability at as high as 100 μM (Fig.2A). It showed that the cell viability exhibited a significant decrease in pretreated cells compared to that in non-pretreated cells (P < 0.05, Fig.2A). The data suggested that pretreatment with LPS increased the cytotoxicity of clopidogrel in HepG2 cells. The results were consistent with the observation that hydrolysis of clopidogrel represents detoxication in overexpression of HCE1 (Tang et al., 2006). However, irinotecan is an anticancer prodrug, and only the hydrolytic metabolite has anticancer activity (Wu et al., 2002). Irinotecan decreased cell viability significantly at the concentration of as high as 30 μM in pretreated cells, while as low as 0.3 μM in the non-pretreated cells (Fig.2B). And the cell viability in pretreated cells exhibited a significant increase compared to that in non-pretreated cells (P < 0.05, Fig.2B). The data suggested that pretreatment with LPS diminished the cytotoxicity of irinotecan in HepG2 cells.

Additionally, the morphological change consisted with the cell viability. As shown in Fig.3, under bright field, cells pretreated with LPS were rounded, isolated, and aggregation, while the cells without LPS pretreatment spread, and the projects were well extended when exposed to 100 μM clopidogrel for 48 h (fig.3, top). Conversely, when exposed to 30 μM irinotecan, cells without LPS pretreatment were isolated and shrank, whereas cells pretreated with LPS were morphologically normal (Fig.3,bottom). The observed morphological changes were opposite to those of cells transfected with HCE1 (Shi et al., 2006; Tang et al., 2006) or HCE2 (Wu et al., 2002).

Fig. 3. Morphological analyses.

Fig. 3

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HepG2 cells were seeded into 96-well plates at the density of 5000 cells/well. After an overnight incubation, 5 μg/ml LPS in 1% serum medium was added to half of the wells, and the treatment lasted for 12 h. Thereafter, the cells were washed with DMEM once, and then cells were treated with 100 μM clopidogrel, or 30 μM irinotecan in 1% serum medium for 48 h (the medium was replaced with fresh drug-containing medium at 24 h). And the images were taken under bright field (200×).

LPS represses HCE1 and HCE2 promoters and decreases the expression of HCE1 and HCE2 through p38MAPK and NF-κB -dependent signaling pathways

To test whether LPS decreased the expression of HCE1 and HCE2 by repressing their promoters, cotransfection was performed with a HCE1 or HCE2 promoter reporter. The HCE1 reporter contains the genomic sequence corresponding to HCE1A2 promoter (HCE1-9.2kb-Luc). As shown in Fig.4A and 4B, treatment with LPS resulted in the decreased activities of HCE1 and HCE2 reporters by 50% and 40% respectively. In order to investigate the mechanisms of the decrease of HCE1 and HCE2 expression mediated by LPS, the differentiated inhibitors, including PDTC (10 μM), SB203580 (5 μM) and U0126 (5 μM) were used 30 min before being treated with LPS in HepG2 cells. As shown in Fig.4, PDTC and SB203580 abolished the decreases of HCE1 and HCE2 mediated by LPS, but U0126 did not. The data suggested that LPS repressed the HCE1 and HCE2 expression through p38MAPK and NF-κB pathways but not through ERK1/2 pathway.

Fig. 4. Transcriptional involvement in HCE1 and HCE2 suppressed by LPS.

Fig. 4

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(A) Repression of HCE1 promoter by LPS. (B) Repression of HCE2 promoter by LPS. HepG2 cells were cultured in 48-well plates at the density of 8×104 and the cells were transfected with HCE1-9.2-Luc (50 ng), or HCE2-Luc (50 ng) along with 5 ng of Null-Renilla reniformis luciferase plasmid. After a 12 h incubation, cells were treated with LPS (5 μg/ml) or the same volume of PBS for another 24 h, then cells were collected and analyzed for luciferase activity. The data are expressed as relative luciferase activity. (C) The pathway of repression of HCE1 and HCE2 mediated by LPS. HepG2 cells were seeded at the density of 5×106 in 6 well-plate overnight and the cells were administrated with PDTC (10 μM), SB203580 (5 μM), or U0126 (5 μM) 30 min before being treated with LPS (5 μg/ml) or the same volume of PBS for 48 h. Cell lysates were prepared and lysates (20 μg) were subjected to Western analyses with an antibody against HCE1, HCE2 or GAPDH. All the experiments were repeated at least three times, and the data were expressed as mean ± SD. * p<0.05, a statistically significant decrease by LPS treatment.

LPS down-regulates mouse carbolesterase 1 and 2 and reduces hydrolytic activity in mice

In order to further confirm that LPS can repress the carboxylesterase 1 and 2 in vivo, male mice were injected intraperitoneally with LPS (5 mg/kg) in saline (Sachdevak et al.,2003), and the controlled mice were treated with the same volume of saline only. After 24 h injection, mice were killed under over anesthetic. The liver and intestine were divided into two parts, with one part being immediately used for preparing total RNA and analyzed by qRT-PCR for mouse carboxylesterase 1 (mCE1) and 2 (mCE2) mRNA levels, and the other one for preparing S9 fractions for the mCE1 and mCE2 protein levels and hydrolysis activity. As shown in Fig.5, LPS reduced the mCE1 and mCE2 of liver at mRNA level by 95% and 90% respectively (Fig.5A), at protein level both by about 85% (Fig.5C), and decreased the hydrolysis activity of liver by 50% (Fig.5E). Meanwhile, LPS also reduced the mCE1 and mCE2 of intestine at mRNA level by 75% and 85% respectively (Fig.5B), at protein level by 40% and 50% respectively (Fig.5D), and decreased the hydrolysis activity of intestine by 50% (Fig.5F). The data suggested LPS also reduced the carboxylesterase 1 and 2 expression and hydrolytic activity in vivo.

Fig. 5. LPS down-regulates mouse carbolesterase 1 and 2 and reduces hydrolytic activity in mice.

Fig. 5

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(A) Effect of LPS on the mCE1 and mCE2 mRNA levels in mouse liver. (B) Effect of LPS on the mCE1 and mCE2 mRNA levels in mouse intestine. (C) Effect of LPS on the mCE1 and mCE2 protein levels in mouse liver. (D) Effect of LPS on the mCE1 and mCE2 protein levels in mouse intestine. (E) Effect of LPS on the hydrolytic activity in mouse liver. (F) Effect of LPS on the hydrolytic activity in mouse intestine. Male mice were injected intraperitoneally with LPS (5 mg/kg) in saline, and control mice were treated with the same volume of saline only. At 24 h after injection for sacrifice, mice were i.p. injected with ketamine (1 ml/kg at 100 mg/ml). After the mice were completely anesthetized (~5 min), the liver was perfused with PBS (37°C) through the portal vein to remove blood. The upper intestine of mice was isolated and washed with cold PBS. The perfused liver and intestine were then divided into two parts, respectively. One part was immediately used for preparing the total RNA and subjected to qRT-PCR analysis for the mRNA levels of mCE1 and mCE2 as described under Materials and Methods. The qPCR Cts were 22 for mCE1, 23 for mCE2 and 18 for mGAPDH. The remaining part was for preparing S9 fractions as described under Materials and Methods for Western analyses and hydrolytic activity assay. All the experiments were repeated at least three times, and the data were expressed as mean ± SD. * p<0.05, a statistically significant decrease by LPS treatment.

Discussion

Hydrolysis of many drugs is reduced in liver diseases such as hepatitis and cirrhosis (Thiollet et al., 1992; Gross et al., 1993). In this study, we have provided a molecular explanation linking LPS induction directly to the decreased capacity of hydrolytic biotransformation. In HepG2 cells, treatment with LPS causes significant decrease in the HCE1 and HCE2 expression, the two major carboxylesterases in the liver. The decreased expression occurs at both mRNA and protein levels, and it is confirmed by enzymatic assay (Fig.1). In the reporter assay, the transcriptional activities of both HCE1 and HCE2 promoters are markedly repressed by LPS (Fig.4A,4B), which suggest that the transcription is involved in the decrease of HCE1 and HCE2 expression mediated by LPS.

In addition, LPS pretreatment profoundly alters the cellular responsiveness to ester drugs, including clopidogrel and irinotecan. The alteration is clearly due to the decrease of the HCE1 or HCE2 expression. More importantly, pretreatment with LPS alters the cellular responsiveness to clopidogrel and irinotecan in an opposite manner to the overexpression of HCE1 and HCE2. For instance, the transfection of HCE1 decreases clopidogrel induced cytotoxicity (Tang et al., 2006), which implies that clopidogrel (parent drug) is much more cytotoxic than its metabolite, whereas pretreatment with LPS increases the cytotoxicity of clopidogrel, which is due to the decrease of HCE1 (Fig.2A, Fig.3top). Likewise, hydrolysis of irinotecan by HCE2 leads to producing SN-38, which is much more cytotoxic to cancer cells (Wu et al., 2002; Charasson et al., 2004; Yang et al., 2007), and the pretreatment with LPS can protect against the cytotoxicity of irinotecan by decreasing HCE2 (Fig.2B, 3 bottom). It should be noted that all ester drugs are tested at submicromolar or micromolar concentrations, and the majority of these concentrations are pharmacologically relevant. Clopidogrel undergoes hydrolytic and oxidative metabolism. The hydrolysis is catalyzed by HCE1 (Tang et al., 2006), whereas the oxidation is catalyzed by CYP3A4 and CYP3A5 (Savi et al., 2000). More importantly, the oxidized but not hydrolytic metabolite exerts antithrombogenic activity, and the hydrolysis of the ester bond represents inactivation (Savi et al., 2000). It is reported that LPS can repress CYP3A4 in human hepatocytes (Aitken and Morgan, 2007). Therefore, the ultimate pharmacological effect probably depends on the relative magnitude of the decreased expression between HCE1 and CYP3A4, with the relative rate between these two types of metabolism. Likewise, irinotecan is an ester prodrug, and the hydrolytic metabolite exerts potent anticancer activity (Masuda et al., 1996; Charasson et al., 2004). In contrast to clopidogrel, irinotecan is hydrolyzed by HCE2 but not HCE1 (Wu et al., 2002). Although the liver expresses high level of HCE2, it has been suggested that hydrolytic metabolite generated locally contributes significantly to its antitumor activity (Rowinsky et al., 1994; Kojima et al., 1998). In support of this notion, gallbladder carcinoma expresses little HCE2, and it represents a natively resistance to irinotecan (Alberts et al., 2002). Our previous study has demonstrated that IL-6 markedly decreases the expression of HCE1 and HCE2 in hepatocytes and hepatoma cells (Yang et al., 2007). In this study, we have also found LPS, which released the proinflammatory cytokines including IL-6, TNFα (Elferink et al., 2004), significantly reduces the HCE1 and HCE2 expression and hydrolytic activity in vitro and in vivo (Fig.1 and 5). Therefore, pretreated with LPS, the cytotoxicity of clopidogrel to HepG2 cells increases, while that of irinotecan decreases (Fig.2, 3).

In order to investigate the mechanism for the decrease of HCE1 and HCE2 expression mediated by LPS, the different selected signal pathway protein inhibitors are used. The results show that both PDTC, a NF-κB inhibitor (Mazzone et al., 2009; Tang et al., 2010), and SB203580, a p38MAPK inhibitor (cheng et al., 2010; Lin et al., 2010), almost abolish the repression of HCE1 and HCE2 protein levels mediated by LPS, but U0126, a selective ERK1/2 inhibitor (Lee and Lee, 2010), does not (Fig.4C). The data suggest LPS down-regulates HCE1 and HCE1 through the p38MAPK and NF-κB pathway but not through the ERK1/2 pathway. Current studies show that inflammatory conditions can activate the p38 mitogen-activated protein kinase (MAPK) signal transduction pathway. That means the phosphorylated, hence activated form of p38, p-p38, regulates the activity of several factors including nuclear factor, NF-κB (Pomerantz and Baltimore, 2002; Tergaonkar, 2006; Papachristou et al., 2008). It has been reported that NF-κB can form a heterdimmer with retinoid X receptor-α (RXR-α). The formation of NF-κB RXR-α heterodimer consumes the RXR-α to disrupt the combination of PXR and RXR-α and therefore represses the CYP3A4 transcription (Gu et al., 2006). Other studies have reported that the nuclear receptors such as PXR and CAR have played a key role in regulating HCE1 and HCE2 in vivo and in vitro (Xu et al., 2009; Raynal et al., 2010; Staudinger et al., 2010). Together, the repression of HCE1 and HCE1 by LPS is likely to be through activating the p38MAPK - NF-κB pathway, and then the activated NF-κB disrupts the function of PXR in the regulation of drug metabolism enzymes including carboxylesterases.

In summary, our work points to several important findings. First, we have shown that LPS causes marked decreases in the expression of HCE1 and HCE2, suggesting that inflammation exerts a broad suppression on the expression of various types of drug-metabolizing enzymes. Second, the transcription was involved in the decrease of HCE1 and HCE2 expression mediated by LPS and the suppression is through p38MAPK - NF-κB pathway. Finally, pretreatment with LPS profoundly alters the cellular responsiveness to ester drugs such as clopidogrel and irinotecan, suggesting that the suppressed expression of carboxylesterases has profound pharmacological and toxicological consequences, particularly with those drugs that are hydrolyzed in an isoform specific manner. Taken together, this study provides new insight into the understanding of the pharmacological and toxicological effects and mechanisms for repressing drug metabolism enzymes in inflammation.

Acknowledgement

The autherors thank Dr. M.E. Dolan for providing plasmid construct. This work is supported by the Natural Science Foundation of China (No. 30772616), the National Key Basic Research Program of China (No.2009CB521906), and the National Institutes of Health grant (USA, F05AT003019, R01ES07965).

Abbreviations

LPS

lipopolysaccharide

NS

normal saline

HCE1

human carboxylesterases1

HCE2

human carboxylesterases2

mCE1

mouse carboxylesterases1

mCE2

mouse carboxylesterases2

CYP450

cytochrome P450

DMEs

drug metabolism enzymes

PXR

pregnane X receptor

hPXR

human pregnane X receptor

TNF-α

tumor necrosis factor α

GAPDH

glyceraldehyde-3-phosphate dehydrogenase

PDTC

ammonium pyrrolidinedithiocarbamate

SB203580

4-(4-Fluorophenyl)-2-(4-methylsulfinylphenyl)-5-(4-pyridyl)-1H-imidazole

U0126

1,4-Diamino-2,3-dicyano-1,4-bis(o-aminophenylmercapto)butadiene monoethanolate

DMSO

dimethysulfoxide

PCR

polymerase chain reaction

DMEM

Dulbecco's modified Eagle's medium

PBS

phosphate-buffered saline

qRT-PCR

quantitative reverse transcription-polymerase chain reaction

p38MAPK

p38 mitogen-activated protein kinase

NF-κB

nuclear factor kappa B

ERK1/2

extracellular regulated kinase1/2

Footnotes

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Conflict of interest statement

The authors declare that there are no conflicts of interest.

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