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Am J Physiol Gastrointest Liver Physiol 290: G377-G385, 2006. First published October 13, 2005; doi:10.1152/ajpgi.00147.2005
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On the role and fate of LPS-dephosphorylating activity in the rat liver

Annemarie Tuin, Ali Huizinga-Van der Vlag, Anne-Miek M. A. van Loenen-Weemaes, Dirk K. F. Meijer, and Klaas Poelstra

Department of Pharmacokinetics and Drug Delivery, Groningen University Institute for Drug Exploration, University of Groningen, Groningen, The Netherlands

Submitted 27 March 2005 ; accepted in final form 22 September 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Gut-derived lipopolysaccharide (LPS) plays a role in the pathogenesis of liver diseases like fibrosis. The enzyme alkaline phosphatase (AP) is present in, among others, the intestinal wall and liver and has been previously shown to dephosphorylate LPS. Therefore, we investigated the effect of LPS on hepatic AP expression and the effect of AP on LPS-induced hepatocyte responses. LPS-dephosphorylating activity was expressed at the hepatocyte canalicular membrane in normal and fibrotic animals. In addition to this, fibrotic animals also displayed high LPS-dephosphorylating activity around bile ducts. The enzyme was shown to dephosphorylate LPS from several bacterial species. LPS itself rapidly enhanced the intrahepatic mRNA levels for this enzyme within 2 h by a factor of seven. Furthermore, in vitro and in vivo studies showed that exogenous intestinal AP quickly bound to the asialoglycoprotein receptor on hepatocytes. This intestinal isoform significantly attenuated LPS-induced hepatic tumor necrosis factor-{alpha} and nitric oxide (nitrite and nitrate) responses in vitro. The enzyme also reduced LPS-induced hepatic glycogenolysis in vivo. This study shows that LPS enhances AP expression in hepatocytes and that intestinal AP is rapidly taken up by these same cells, leading to an attenuation of LPS-induced responses in vivo. Gut-derived LPS-dephosphorylating activity or enzyme upregulation within hepatocytes by LPS may therefore be a protective mechanism within the liver.

lipopolysaccharide; inflammation; intestine; alkaline phosphatase

LIPOPOLYSACCHARIDE (LPS) or endotoxin is a highly toxic molecule derived from the outer membrane of gram-negative bacteria. The toxic part of LPS is located in the lipid A part of the molecule, which is the most well conserved moiety among LPS serotypes from a variety of gram-negative bacteria. Two phosphate groups coupled to two glucosamines in the lipid A part largely determine the toxicity of LPS (17, 39). Removal of a single phosphate group results in the formation of a nontoxic monophosphoryl lipid A moiety (6, 24, 46).

Systemic LPS can elicit a systemic inflammatory response syndrome characterized by fever, diffuse intravascular coagulation, shock, multiple organ failure, and eventually death (2, 16). At present, sepsis is still the most important cause of death in intensive care units (25) and the 10th cause of death overall in the United States (5), accounting for about 215,000 deaths/yr (4). Elevated LPS levels in the blood may be the result of a serious bacterial infection, an impaired host defense system, or damage to barriers like the skin or the intestine or, alternatively, from a reduced capacity of the liver to remove LPS from the blood. Several diseases are now associated with bacterial translocation from the gut (42a), e.g., trauma, hemorrhagic shock, burn injuries, inflammatory bowel disease, cardiovascular and liver surgeries, acute pancreatitis, and necrotizing enterocolitis. Impaired clearance of LPS by the liver is usually observed during liver diseases like cirrhosis and hepatitis B and C (22, 43). These diseases demonstrate the importance of an intact liver and intestinal wall to prevent LPS-induced diseases.

The liver is the major LPS-removing organ. The majority of systemic LPS is removed from the bloodstream by Kupffer cells and likely also endothelial cells. Kupffer cells modify the endocytosed LPS (48) and somehow pass it on to hepatocytes, which subsequently excrete these products into the bile (18). Part of the LPS, though, is removed from the circulation directly by hepatocytes (40). Direct uptake of LPS by hepatocytes is significantly enhanced by binding of LPS to proteins such as apolipoprotein E (38) and particles like chylomicrons (12, 13), very-low-density lipoprotein, low-density lipoprotein (11), and high-density lipoprotein (11).

Poelstra et al. (33, 34) were the first to postulate a role of the endogenous enzyme alkaline phosphatase (AP) in LPS detoxification. This enzyme appeared to be able to remove phosphate groups from LPS, thus attenuating the toxicity of this molecule. Until then, the only physiological role for AP that had been identified was a role in bone formation (36).

Because the enzyme is present within the liver, and in fact is upregulated during several liver diseases, we now addressed the question of whether this enzyme plays a role in the protection of the liver against LPS. Cholestasis and sepsis are characterized by both elevated serum AP and LPS levels (23, 49). Commonly, these elevated AP levels are regarded as a reflection of liver damage. In view of the LPS-detoxifying activity of AP, we now tested whether these elevated AP levels may be an adaptive physiological response upon LPS. To answer this question, we examined the LPS-dephosphorylating activity of the liver and measured intrahepatic changes in this activity after LPS administration and after the induction of fibrosis both at the enzyme and RNA level.

In addition, AP activity is also high on the membranes of enterocytes lining the gut, and this intestinal enzyme is also present as a soluble enzyme in plasma, from where it is rapidly taken up by hepatocytes (3, 7, 26). This expression pattern and pharmacokinetic profile might be indicative for a hepatoprotective role. Therefore, we also tested whether exogenous intestinal AP (iAP) might protect hepatocytes from LPS-induced damage. Our data show that iAP is likely to be targeted to hepatocytes and support the hypothesis that this enzyme is part of the endogenous defense system against LPS.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chemicals. Highly purified calf iAP (ciAP; specific activity: 2,530 U/mg) was obtained from AM-Pharma (Bunnik, The Netherlands). LPS from Escherichia coli (serotype O55:B5), obtained from Sigma (St. Louis, MO), was used in all in vitro and in vivo studies. All other chemicals were of analytical grade.

Animals. Male Wistar rats (Harlan; Zeist, The Netherlands) were housed under standard laboratory conditions under a regular light-dark cycle with laboratory chow and acidified water ad libitum. All animal experiments were approved by the Local Committee on Animal Experimentation.

Enzyme histochemistry. LPS-dephosphorylating activity of endogenous AP in rat livers was examined by incubating cryostat sections (5 µm) with LPS as a substrate according to Wachstein and Meisel (27). Briefly, sections were fixed in 4% formalin-macrodex and subsequently incubated for 120 min in Tris·HCl buffer (pH 7.6) containing LPS (final concentration: 3.2 mg/ml), MgSO4 (final concentration: 0.01 M) and Pb(NO3)2 [final concentration: 0.06% (wt/vol)] at 37°C. A lead phosphate precipitate is formed at the site of enzyme activity, which is converted by incubation with Na2S to a lead sulphate precipitate, which appears as a dark brown staining. Specificity of this staining has previously been demonstrated by inhibition of AP activity using levamisole (34) and L-phenylalanine (47). LPS was omitted in control incubations.

Immunohistochemical staining of the CD14 receptor. Acetone-fixed cryostat sections (5 µm) of the rat liver, lung, kidney, and small intestine from rats with established liver fibrosis 3 wk after bile duct ligation (BDL3) were prepared and subsequently incubated with a rabbit polyclonal antibody against the CD14 receptor (CD14r; cat. no. sc-9150, Santa Cruz Biotechnology) or PBS (control sections). After inhibition of endogenous peroxidase (0.25% H2O2), sections were incubated with goat anti-rabbit IgG conjugated with horseradish peroxidase. Peroxidase activity was visualized with 3-amino-9-ethylcarbazole according to standard methods. Finally, sections were counterstained with hematoxylin.

Immunohistochemical staining of collagen III. This staining was performed as described for the CD14r staining. A goat polyclonal antibody against collagen III (cat. no. 1330-01, Southern Biotechnology Associates; Birmingham, USA) was used, and rabbit anti-goat IgG conjugated with horseradish peroxidase was used as a secondary antibody.

In vivo experiments. Experiments were carried out in normal rats and BDL3 rats. The rats were divided into three groups. Control animals received saline intravenously at time (t) = 0, whereas others received 1 mg/kg LPS intravenously at t = 0. Some animals were treated with 100 or 1,000 units ciAP intravenously 1 min before the LPS injection. After 2 h, rats were killed under O2-N2O-forene (Isoflurane, Abbott Laboratories; Kent, UK) anaesthesia. Livers were taken out, partly frozen in isopentane, stored at –80°C for immunohistochemical staining, and partly snap frozen in liquid nitrogen for RNA isolation. Blood was collected to measure liver enzymes. If indicated, kidneys, lungs, and intestines were taken out and handled as described above for the liver.

Slice experiments. BDL3 rats were killed under O2-N2O-forene anaesthesia, and livers were taken out and stored in University of Wisconsin organ preservation solution (UW solution) until slice preparation. Precision-cut liver slices (10–14 mg) were prepared as described previously (29) and stored in UW solution on ice until incubation. Slices were incubated individually at 37°C in six-well plates (Greiner) in 3.2 ml Williams' medium E supplemented with glutamax I (GIBCO-BRL; Paisley, Scotland) and 50 mg/ml gentamicin (GIBCO-BRL) and saturated with 95% O2-5% CO2. Slices were incubated for 24 h with or without 10 µg/ml LPS. Slices incubated with LPS were incubated with or without ciAP (1.56 or 15.6 U/ml). Medium of the slices was stored at –20°C until nitric oxide [nitrite and nitrate (NOx)] measurements or at –80°C until TNF-{alpha} measurements. Also, slices were embedded in Tissue-Tek and stored at –80°C until the preparation of cryostat sections.

Real-time quantitative PCR analysis. RNA was isolated from rat liver pieces using the Qiagen RNeasy Mini Kit. Quality of the RNA was checked on a 2% agarose gel, and the RNA concentration was determined with a ribogreen assay. The Reverse Transcription System from Promega was used to convert 1.6 µg RNA into cDNA. RT-PCR was performed for 10 s at 25°C, 60 s at 45°C, and 5 s at 95°C using olido-dT (0.633 µg/reaction) primers. Then, 1.25 µl cDNA was amplified by quantitative real-time PCR using the SYBR green PCR Master Mix from Applied Biosystems. PCRs were carried out in a ABI PRISM 7900HT Sequence Detection System (Applied Biosystems) in a volume of 21.25 µl. Genes of interest were amplified using the following primers: GAPDH, forward 5'-CCATCACCATCTTCCAGGAG-3' and reverse 5'-CCTGCTTCACCACCTTCTTG-3'; liver/bone/kidney AP (LBK-AP), forward 5'-GCAAGGACATCGCCTATCAG-3' and reverse 5'-AGTTCAGTGCGGTTCCAGAC-3', and glutamate-pyruvate transaminase (GPT), forward 5'-TGTGCCTCCTGGAAGAGACT-3' and reverse 5'-TGTTGCGTCAGAGACTGTCC-3'. GAPDH was used as a housekeeping gene, and levels of AP and GPT were normalized to GAPDH levels. Data obtained were analyzed using the comparative threshold cycle (CT) method as described in User Bulletin No. 2 of the ABI PRISM 7700 Sequence Detection System (Applied Biosystems). Data are expressed as fold induction or repression of the gene of interest compared with the control condition as calculated by the formula 2–{Delta}{Delta}CT.

NOx assay. NOx production by rat liver slices was assayed by determining NOx levels in the media of slices. This assay was performed according to Bentala et al. (6).

TNF-{alpha} assay. TNF-{alpha} levels in rat serum and media of liver slices were determined using a sandwich ELISA according to Bentala et al. (6). Ninty-six-well ELISA plates were coated overnight with a monoclonal anti-rat TNF-{alpha} capturing antibody (R&D Systems). Sera from rats were diluted two or five times before they were tranferred to the plate, whereas samples from the incubation media of rat liver slices were measured undiluted. Recombinant-rat TNF-{alpha} (R&D Systems) was used to prepare a TNF-{alpha} standard curve. Biotinylated rabbit anti-mouse/rat TNF-{alpha} (R&D Systems) was used as a detection antibody according to standard methods.

Periodic acid Schiff staining. The glycogen content of hepatocytes was visualised by periodic acid Schiff (PAS) staining. Briefly, cryostat sections (5 µm) were fixed in 4% (vol/vol) formalin in methanol, subsequently incubated in 1% periodic acid, washed, incubated in Schiff's reagents, washed with tap water, and counterstained with hematoxylin. The hepatic glycogen content appears as purple staining.

Clearance of iAP and uptake by hepatocytes. Uptake of exogenous AP by hepatocytes was studied in rats in vitro and in vivo. In vivo, three rats received 100 units ciAP intravenously, and three rats received 10 mg asialofetuin (AsF) intravenously 1 min before the ciAP injection. Blood samples were taken at t = 0, 1, 2, 5, 10, 20, 30, and 60 min after ciAP administration. Serum AP activity was measured at pH 9.8 using para-nitrophenyl phosphate as a substrate according to standard methods (9). Pharmacokinetic parameters were assessed with the computer program MultiFit (Dr. J. H. Proost; Groningen, The Netherlands).

In vitro, precision-cut rat liver slices were incubated as described above for 2 h with 15.6 U/ml ciAP alone or with 15.6 U/ml ciAP plus 0.5 mg/ml AsF, and 5-µm cryostat sections were stained for AP activity using {beta}-glycerophosphate as a substrate according to the method of Gomorri (21).

Statistics. Results are expressed as means ± SD of at least three separate experiments. Data were analyzed using a two-sided Student's t-test and considered significant at P < 0.05.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
LPS dephosphorylation in the rat liver. Because the liver is the major LPS-removing organ of the body, we first tested whether the liver itself is capable of dephosphorylating, and thus detoxifying, LPS by means of its endogenous AP activity. The LPS-dephosphorylating activity of rat livers was examined in 5-µm cryostat sections using an enzymehistochemical staining method at pH 7.6 with LPS as a substrate and lead phosphate as the product. LPS-dephosphorylating activity was examined in normal as well as BDL3 rat livers, which are known to have elevated AP levels. Figure 1 shows LPS dephosphorylation in cryostat sections of normal (A) and BDL3 (B and C) rat livers. Control sections (Fig. 1D), which were incubated with vehicle instead of LPS, showed no staining at all. In normal rat livers, a clear dark brown staining, reflecting the dephosphorylating activity, was found. Lead phosphate precipitates were located around hepatocytes, showing a clear canalicular staining pattern (8). In BDL3 rats, a similar staining pattern was found around hepatocytes, but, in addition to this, very strong staining around bile ducts was also seen (Fig. 1C).



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  		Fig. 1. Staining for alkaline phosphatase (AP) activity in the rat liver using lipopolysaccharide (LPS) as a substrate. Dark staining (arrows) indicates sites of LPS dephosphorylation (PbS precipitate). Enzyme activity in representative liver sections from normal rats (A) and from rats with established liver fibrosis 3 wk after bile duct ligation (BDL3; B and C) are shown. D: control rat liver section incubated without the substrate LPS (magnification: x400). Sections of normal and BDL3 rat livers were also stained for collagen III deposition to show that fibrosis had developed 3 wk after bile duct ligation. The collagen III-stained sections are shown as insets in A and B (magnification: x100). The inset in C depicts a higher magnification of bile duct epithelial (BDE) cells (magnification: x1,000). The inset clearly shows that AP activity is present along BDE cells (black staining) but that these cells themselves do not show any AP activity.

 
Dephosphorylation of different LPS serotypes by endogenous hepatic AP. To investigate whether rat liver AP is capable of dephosphorylating LPS from several bacterial species, enzymehistochemical staining was also carried out with LPS from five other bacterial species: in addition to wild-type E. coli, we tested Salmonella typhimurium, Pseudomonas aeruginosa (serotype 10), Shigella flexneri (serotype 1A), Klebsiella pneumonia (Sigma), and Salmonella minnesota Re 595 (List Biological Laboratories; Campbell, CA). Light microscopical evaluation clearly showed that LPS from all bacterial species tested was dephosphorylated, although different staining intensities were found (Table 1). A semiquantitative evaluation by two independent observers was performed by ranking the sections in order of increasing staining intensity.


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  		Table 1. Semiquantitative evaluation of LPS-dephosphorylating activity in rat liver sections using LPS from six different bacterial species in BDL3 rat liver sections

 
Localization of CD14r and AP activity in rats. Together with Toll-like receptor 4 (TLR4) and myeloid differentiation protein 2 (MD-2), CD14r is part of the LPS signaling system. Because AP is present at sites where possible pathogens, e.g., gram-negative bacteria, may enter the body (e.g., lung epithelium and the intestine) or where LPS may accumulate (the liver and kidney), it is interesting to see how AP activity is related to CD14r expression. To address this issue, an immunohistochemical staining of CD14r was performed on cryostat sections of BDL3 rat livers, lungs, kidneys, and intestines. This staining was compared with the staining of AP activity in these organs.

In the kidney, we found a striking colocalization of AP activity and CD14r expression. Both AP activity and CD14r were located at the brush borders in renal tubuli (Fig. 2, C and D). Also, in lungs, livers, and intestines, AP activity and CD14r expression were found, although they did not completely colocalize. In lungs, CD14r expression was detected at the apical side of alveolar epithelial cells (Fig. 2E), whereas AP activity was detected at the basal side of alveolar epithelial cells (Fig. 2F). In livers, CD14r expression was localized on bile duct epithelial cells (Fig. 2A), whereas AP activity was detected around the bile duct on fibroblast-like cells (Fig. 2B). In contrast, hepatocytes that showed AP activity along their membrane did not show any detectable CD14r expression. Another study (44), however, has reported low CD14r expression in hepatocytes, whereas a recent study (31) has also indicated CD14r expression in activated human stellate cells (myofibroblasts) around proliferative bile ducts. Within the small intestine, AP activity was found along the entire villus (Fig. 2H), whereas CD14r expression was only detected in the crypts and the lowest part of the villus (Fig. 2G). This corresponds with data in the literature (30). In the small intestine, there is thus a colocalization of AP activity and CD14r expression in the lower half of the villi at the bottom of the crypts. In conclusion, AP and CD14r were localized in all organs tested, with a colocalization in the kidneys, liver, and intestine. Within the lung, both are in close proximity of each other. An exception seems to be the arterial wall. In the tunica adventitia of blood vessels, high AP activity was found (Fig. 2D), but no CD14r expression was seen at all.



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  		Fig. 2. CD14 receptor (CD14r) expression and AP activity in the rat liver (A and B), kidney (C and D), lung (E and F), and small intestine (G and H). A, C, E, and G show the localization of CD14r (red), whereas B, D, F, and H show AP activity (brown). There is occasional staining for CD14r in all organs examined. Staining for AP activity is also seen in every organ. AP staining is frequently localized in the same areas as CD14r staining (for details, see the text). I and J: negative controls (omission of the first antibody) of the two organs in which CD14r staining was most intense, that is, the liver (I) and kidney (J). a, Artery. Magnification: x400.

 
AP activity after LPS administration in vivo. From the literature, it is known that several diseases are associated with elevated AP levels as well as high LPS levels, e.g., cholestasis, sepsis, and liver fibrosis. To test whether both events can be related to each other, rats received LPS (1 mg/kg) or saline intravenously, and the effect on AP activity in the blood and AP mRNA levels in livers was assayed. We studied mRNA levels 2 h after the injection of LPS to exclude a role for the many mediators that are produced several hours after a LPS challenge. After 2 h, normal rats did not show a significant change in serum AP levels in response to LPS administration (Fig. 3A). In the literature, increases of AP activity in serum after LPS administration were also reported in normal rats but in other conditions, e.g., at intervals longer than 2 h after LPS injection and at higher LPS concentrations (19). Also, the levels of other markers of liver damage (37), lactate dehydrogenase (LDH), glutamate-oxalacetate transaminase (GOT), GPT, and {gamma}-glutamyl transferase did not show a significant elevation 2 h after the LPS challenge (data not shown). In contrast, in serum of BDL3 rats, significant increases in serum markers were found at t = 2 h (Fig. 3A). AP activity in serum rose from 414 ± 48 U/l in saline-treated rats to 541 ± 94 U/l in LPS-treated rats (P < 0.05). This paralleled an increase in serum levels of LDH (from 2,674 ± 1,581 to 5,835 ± 2,114 U/l), GOT (from 374 ± 116 to 1,466 ± 483 U/l), and GPT (from 78 ± 15 to 134 ± 23 U/l).



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  		Fig. 3. Serum AP levels and intrahepatic mRNA levels for AP and glumatate-pyruvate transaminase (GPT) in normal and BDL3 rats 2 h after the administration of saline or LPS. A: serum AP levels. B: intrahepatic mRNA expression for AP. C: intrahepatic mRNA expression for GPT. It can be seen that LPS causes an elevation in serum AP levels in BDL3 rats but not in normal rats. In contrast, mRNA levels for AP are significantly elevated in both normal and BDL3 livers at this time point. These elevated AP mRNA levels are not associated with increased intrahepatic GPT mRNA levels (C). Values are means ± SD; n = 4 rats/group. *P < 0.05 vs. saline.

 
To examine whether the elevated AP levels are the result of release from damaged hepatocytes or de novo synthesis, we studied mRNA expression in these livers using real-time PCR. In both normal and BDL3 rats, a significant induction of AP mRNA could be detected 2 h after LPS administration (Fig. 3B). Real-time PCR revealed a sevenfold induction of AP mRNA in normal livers of rats receiving LPS compared with control rats receiving saline. BDL3 rats showed almost the same result but with a maximum fold of induction of five. This increase could be mimicked in vitro by incubation of liver slices with LPS. In these slices, there is no influx of neutrophils (which have high mRNA levels for AP), thus excluding a role for these cells as the source of this mRNA (data not shown).

To assess whether this induction in mRNA levels for AP parallels the increase in markers reflecting liver damage, we also measured GPT mRNA levels. Figure 3B depicts GPT mRNA levels, and the data clearly show that these mRNA levels remained constant (normal rats) or even slightly decreased (BLD3 rats). This means that, although both serum GOT and serum AP are elevated after LPS in BDL3 rats, only in the case of AP is this associated with an increase in mRNA for AP. Elevated AP and GPT levels in the blood observed after LPS-induced liver damage therefore seem to have a different physiological background.

Uptake of iAP by hepatocytes. iAP is a glycoprotein with terminal galactose groups that is produced by enterocytes. From the literature, it is known that iAP originating from dogs is taken up from the circulation by the asialoglycoprotein receptor (ASGP-R) on hepatocytes (7, 26). We also tested this in our system with our highly purified ciAP preparations. To this end, we examined the plasma half-life of ciAP in serum and also tested the effect of AsF, a well-known blocker of ASGP-R, on plasma disappearance. Our results (Fig. 4A) clearly confirm the particular literature data. The plasma concentration versus time curve in rats that only received ciAP showed a short distribution half-life of 2.8 min and an elimination half-life of 39 min. Assessment of the effect of AsF on the pharmacokinetic profile showed that AsF can almost completely prevent ciAP clearance from the blood. These data demonstrate that ASGP-R is most likely involved in the clearance of ciAP.



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  		Fig. 4. A: serum AP activity after an intravenous administration of 100 U calf intestinal AP (ciAP) with or without prior administration of the asialoglyoprotein receptor blocker asialofetuin (AsF; 10 mg/rat). Serum AP activity was measured at pH 9.8 at different time points after ciAP administration. B: AP activity in rat liver slices. Rat liver slices were incubated for 2 h with 15.6 U/ml ciAP alone (top) or with 15.6 U/ml ciAP plus 0.5 mg/ml AsF (bottom) and thereafter embedded in Tissue-Tek. Cryostat sections were stained for AP activity at pH 9 using {beta}-glycerophosphate as a substrate according to the method of Gomorri (21). Arrows indicate the presence of AP activity in hepatocytes. It can be seen that AsF blocks the uptake of ciAP in vivo and in vitro. Magnification: x400.

 
We also examined binding and uptake of iAP by hepatocytes in vitro. To show these events, rat liver slices were incubated with ciAP alone or with ciAP plus AsF, and AP activity was detected by enzymehistochemistry (Fig. 4B). In slices incubated with ciAP alone, high AP activity was found in the cytoplasm of hepatocytes, whereas in slices incubated with ciAP plus AsF, no AP activity was detected at all. The absence of AP activity in hepatocytes treated with ciAP plus AsF confirmed that ciAP is preferentially bound to and taken up via ASGP-R on hepatocytes.

In vitro studies with iAP. To assess the effect of exogenous iAP on hepatocytes in vitro, slices prepared from BDL3 rat livers were incubated with 10 µg/ml LPS for 24 h with or without ciAP. Concentrations of 1.56 and 15.6 U/ml ciAP were tested, and NOx production in the medium was assayed. After stimulation of BDL3 liver slices with LPS, NOx levels in the medium rose from 5.4 ± 1.3 µM (control without LPS) to 114.4 ± 13.7 µM at t = 24 h after the addition of LPS. Importantly, our data also showed that NOx levels in the medium were significantly reduced when 1.56 and 15.6 U/ml ciAP were added to the medium (Fig. 5A).



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  		Fig. 5. Nitric oxide [nitrite and nitrate (NOx)] and TNF-{alpha} levels in the medium of rat liver slices after incubation with LPS and ciAP. A: NOx levels in the medium of rat liver slices in response to LPS. BDL3 rat liver slices were incubated for 24 h with 10 µg/ml LPS with or without 1.56 or 15.6 U/ml ciAP. B: TNF-{alpha} levels in the medium from BDL3 liver slices treated with LPS and incubated with or without ciAP. BDL3 rat liver slices were incubated for 24 h with 10 µg/ml LPS with or without 1.56 or 15.6 U/ml ciAP. TNF-{alpha} and NOx production of liver slices under basal conditions was 46 ± 27 pg/ml and 5.4 ± 1.3 µM, respectively, and, after the addition of LPS, levels rose to 560 ± 108 pg/ml and 114.4 ± 13.7 µM, respectively. These amounts were set to 100% in each experiment. Values are means ± SD of 3 different experiments (3 slices/experiment). *P < 0.05 vs. no ciAP.

 
In addition, we looked at the TNF-{alpha} response in slices upon LPS. For these experiments, only slices from BDL3 rat livers were used, because these rats produced much higher amounts of TNF-{alpha} in response to LPS. After incubation of the slices with LPS, TNF-{alpha} levels reached a maximal response at t = 24 h, and the level had risen from 46 ± 27 pg/ml (control without LPS) to 560 ± 108 pg/ml (data not shown). We observed (Fig. 5B) that slices incubated with ciAP displayed a significantly lower TNF-{alpha} response upon LPS compared with slices incubated with LPS alone (P < 0.05).

In vivo studies with iAP. We also tested the effect of LPS and ciAP on hepatocytes in vivo. Because of the early time point of death after the LPS challenge (t = 2 h), endogenous AP activity was not increased yet (data not shown), and only direct effects of LPS could be found.

One of the earliest effects of LPS on the liver is the stimulation of glycogenolysis (45). This glycogen degradation is an early event in hepatocytes responding to stress or illness. We therefore examined the glycogen content of hepatocytes in rat livers after LPS administration using PAS staining. Figure 6 clearly shows that 2 h after LPS administration, the glycogen content in hepatocytes is very strongly reduced in livers of rats that received LPS (B) compared with rats that received saline (A). All of the rats (n = 9) treated with LPS showed a strong reduction in PAS staining throughout the whole liver (Fig. 6B). In contrast, all of the rats (n = 6) treated with 1,000 units ciAP after LPS clearly exhibited partially retained PAS staining (Fig. 6C), whereas none of the control rats or rats receiving ciAP alone (n = 3) or saline (n = 8) displayed a significant disappearance of PAS staining.



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  		Fig. 6. Glycogen staining in hepatocytes as detected by periodic acid Schiff (PAS) staining on rat liver cryostat sections. Rats received saline (n = 9; A), LPS (1 mg/kg, n = 9; B), or LPS (1 mg/kg) plus ciAP (1,000 units, n = 6; C) intravenously. Note the reduced PAS staining in LPS-treated rats (B) compared with control rats (A), whereas PAS staining was higher in rats that received LPS plus ciAP (C) compared with ciAP-untreated rats (B). Magnification: x100.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Numerous studies have described AP activity in the liver and changes in this activity during diseases. Yet, this was not related to a possible physiological role for this enzyme. Until recently, the only known physiological function of AP was its role in bone formation (36), but, obviously, the enzyme fulfils another function in the intestine, kidney, liver, and along blood vessels. All of these organs are located at sites in close contact with the external environment and represent sites where pathogens, e.g., gram-negative bacteria, may enter the body. The presence of AP at these locations as well as the simultaneous elevation of AP and LPS levels in the blood in several diseases combined with the observation that AP is able to dephosphorylate and detoxify LPS led us to the idea that another physiological function of AP could be the detoxification of LPS (6, 33, 34).

This study, therefore, focused on the role of LBK-AP in the liver and on the role of iAP in LPS dephosphorylation. We found that the liver by means of its AP is able to dephosphorylate LPS and that LPS, in turn, is able to induce LBK-AP mRNA levels. Whether this is a direct effect of LPS or mediated by other factors remains to be elucidated, but the effect occurs very rapidly at a time point when most inflammatory mediators are not yet elevated.

Within the liver, LPS-dephosphorylating activity was present in hepatocytes but also in nonparenchymal cells, most likely fibroblasts and Kupffer cells (Fig. 1). In BDL3 rats, the nonparenchymal staining was mainly visible around the bile ducts and blood vessels. Localization of LPS-dephosphorylating activity corresponded with known AP localization in the liver and was inhibited by the LBK-AP inhibitor levamisole (34). It should be noted that this activity of the "alkaline" enzyme is detectable at pH 7.4. When AP activity was measured with conventional substrates ({beta}-glycerophosphate) in the same concentration as LPS (~160 µM) and at pH 7.4, no dephosphorylating activity was found at all, indicating that at pH 7.4 LPS is a better substrate for the enzyme than {beta}-glycerophosphate. We showed in the present study that LBK-AP was able to dephosphorylate LPS from different bacterial species, albeit to a different extent (Table 1). This may be related to differences in chemical structure between the various LPS molecules that cause differences in solubility and micelle formation or to differences in molecular weight, resulting in differences in molarities of solutions with the different LPS molecules. The molecular weight of wild-type LPS molecules cannot be determined accurately (32).

LPS circulating in the plasma can activate LPS-responsive cells via LPS-binding protein (LBP). The LBP-LPS complex binds to CD14r (35), which is present in the membrane of, among others, monocytes and macrophages (14). Cells lacking membrane-bound CD14r, like endothelial cells, can be activated via soluble CD14r, present in serum (15). The LBP-LPS-CD14r complex associates with TLR4 and MD-2, resulting in intracellular signaling (1, 42). Epithelial cells that do not express membrane-bound CD14r and MD-2 can be activated by soluble forms of both CD14r and MD-2 (15, 20, 35). Because AP may be involved in LPS detoxification, a distribution pattern of these LPS-capturing molecules similar to AP was anticipated. In the rat kidney, liver, and small intestine, but not in the blood vessel wall, CD14r colocalized with LPS-dephosphorylating activity, whereas in the rat lung, CD14r expression and AP activity were detected in close proximity to each other. The present results support and extend another study (10) that showed a colocalization of AP and CD14 in intracellular compartments in neutrophils and is in line with the newly proposed function of the enzyme.

We also studied the influence of LPS on AP expression in the body and focused on hepatic expression of the LBK-AP isoenzyme. Although AP induction by LPS in cardiac muscle has already been reported (28), we show for the first time that LPS directly induces AP mRNA levels in normal as well as BDL3 rat livers. The increment in hepatic mRNA levels likely precedes the rise in serum AP levels in normal rats (Fig. 3, A and B). Other reports support this hypothesis by showing a rise in serum AP levels at later time points after LPS injection (19). BDL3 rats are characterized by enhanced serum AP levels (Fig. 3A) already at t = 2 h and increased basal mRNA levels for AP compared with normal rats (data not shown). The reason for this increased basal AP expression and increased serum levels after bile duct ligation remains to be elucidated, but, again, this might be a physiological response to the increased LPS levels that occur after bile duct ligation (23, 41).

The next question we addressed was whether AP is able to protect the liver, and especially hepatocytes, from LPS-induced damage. This was examined in vitro in rat liver slices and in vivo in rats using exogenous ciAP. We used exogenous iAP to mimic the induction of endogenous AP on the plasma membrane of hepatocytes. From the literature, it is known that iAP rapidly binds to ASGP-R, which is present on hepatocytes (7, 26). We confirmed this by using a known blocker of ASGP-R, AsF, and studied its effects on the clearance of highly purified ciAP (Fig. 4). In addition, binding and uptake of iAP by hepatocytes was confirmed in vitro in rat liver slices by enzymehistochemical staining for AP activity.

The effect of ciAP on hepatocytes in vitro was assessed in BDL3 liver slices, which contain all the resident liver cell types in their original phenotype and context, including the cell-cell contacts. TNF-{alpha} and NOx were rapidly produced by hepatic cells in response to LPS, and the addition of ciAP to the medium significantly reduced both NOx and TNF-{alpha} levels (Fig. 5, A and B).

Also, in vivo, ciAP exhibited a protective effect. Our hypothesis that iAP protects hepatocytes against LPS-induced damage was supported by glycogen PAS staining on rat livers. We explored the glycogen content of hepatocytes as a parameter because it is strikingly influenced by LPS (45). On the basis of this parameter, our study clearly showed a protective effect of iAP in all rats examined after a challenge with LPS.

In conclusion, our results demonstrate that liver AP dephosphorylates LPS (Fig. 1 and Table 1) and that the LPS-dephosphorylating activity in various organs colocalizes with or is closely located near CD14r (Fig. 2). Furthermore, LPS rapidly induces mRNA for AP in hepatocytes while, at the same time, iAP is rapidly transferred from the intestine to hepatocytes (Fig. 4). The combined AP induction and AP transfer from the small intestine to the liver likely reflects a physiological response to LPS. It is well known that dephosphorylated LPS is nontoxic (6, 24, 46). The hepatoprotective effect of AP is supported by a reduction of TNF-{alpha} and NOx levels in vitro (Fig. 5) and decreased glycogenolysis (Fig. 6) in hepatocytes in vivo induced by iAP. Our study may provide new perspectives for a possible treatment of gram-negative sepsis and illustrates the relevance of cross talk between the liver and intestine.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was financially supported by the Dutch Organization for Scientific Research.


   ACKNOWLEDGMENTS
 
We thank Dr. R. Brands (AM-Pharma; Bunnik, The Netherlands) for the generous supply of ciAP.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. Tuin, Dept. of Pharmacokinetics and Drug Delivery, Univ. of Groningen, Antonius Deusinglaan 1, 9713 AV Groningen, The Netherlands, (e-mail: a.tuin{at}rug.nl)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Aderem A and Ulevitch RJ. Toll-like receptors in the induction of the innate immune response. Nature 406: 782–787, 2000.[CrossRef][Medline]
  2. Aird WC. The role of the endothelium in severe sepsis and multiple organ dysfunction syndrome. Blood 101: 3765–3777, 2003.[Abstract/Free Full Text]
  3. Alpers DH, Zhang Y, and Ahnen DJ. Synthesis and parallel secretion of rat intestinal alkaline phosphatase and a surfactant-like particle protein. Am J Physiol Endocrinol Metab 268: E1205–E1214, 1995.[Abstract/Free Full Text]
  4. Angus DC, Linde-Zwirble WT, Lidicker J, Clermont G, Carcillo J, and Pinsky MR. Epidemiology of severe sepsis in the United States: analysis of incidence, outcome, and associated costs of care. Crit Care Med 29: 1303–1310, 2001.[CrossRef][ISI][Medline]
  5. Arias E, Anderson RN, Kung HC, Murphy SL, and Kochanek KD. Deaths: final data for 2001. Natl Vital Stat Rep 52: 1–115, 2003.[Medline]
  6. Bentala H, Verweij WR, Huizinga-Van der Vlag A, Loenen-Weemaes AM, Meijer DK, and Poelstra K. Removal of phosphate from lipid a as a strategy to detoxify lipopolysaccharide. Shock 18: 561–566, 2002.[CrossRef][ISI][Medline]
  7. Blom E, Ali MM, Mortensen B, and Huseby NE. Elimination of alkaline phosphatases from circulation by the galactose receptor. Different isoforms are cleared at various rates. Clin Chim Acta 270: 125–137, 1998.[CrossRef][ISI][Medline]
  8. Cormack DH. The digestive system. In: Essential Histology. Philadelphia, PA; Lippincott-Raven, 1993, p. 271–306.
  9. Courdouhji MK, Guelfi JF, Fontanilles AM, Grozdea JD, and Vergnes HA. Characterization of alkaline phosphatase in polymorphonuclear neutrophils from normal sheep. Enzyme 40: 217–222, 1988.[ISI][Medline]
  10. Detmers PA, Zhou D, Powell D, Lichenstein H, Kelley M, and Pironkova R. Endotoxin receptors (CD14) are found with CD16 (Fc gamma RIII) in an intracellular compartment of neutrophils that contains alkaline phosphatase. J Immunol 155: 2085–2095, 1995.[Abstract]
  11. Flegel WA, Baumstark MW, Weinstock C, Berg A, and Northoff H. Prevention of endotoxin-induced monokine release by human low- and high-density lipoproteins and by apolipoprotein A-I. Infect Immun 61: 5140–5146, 1993.[Abstract/Free Full Text]
  12. Harris HW, Grunfeld C, Feingold KR, Read TE, Kane JP, Jones AL, Eichbaum EB, Bland GF, and Rapp JH. Chylomicrons alter the fate of endotoxin, decreasing tumor necrosis factor release and preventing death. J Clin Invest 91: 1028–1034, 1993.[ISI][Medline]
  13. Harris HW, Rockey DC, and Chau P. Chylomicrons alter the hepatic distribution and cellular response to endotoxin in rats. Hepatology 27: 1341–1348, 1998.[CrossRef][ISI][Medline]
  14. Haziot A, Chen S, Ferrero E, Low MG, Silber R, and Goyert SM. The monocyte differentiation antigen, CD14, is anchored to the cell membrane by a phosphatidylinositol linkage. J Immunol 141: 547–552, 1988.[Abstract]
  15. Henneke P and Golenbock DT. Innate immune recognition of lipopolysaccharide by endothelial cells. Crit Care Med 30: S207–S213, 2002.[CrossRef][ISI][Medline]
  16. Heumann D and Glauser MP. Pathogenesis of sepsis. Scientific American 1: 28–37, 1994.
  17. Holst O, Thomas-Oates JE, and Brade H. Preparation and structural analysis of oligosaccharide monophosphates obtained from the lipopolysaccharide of recombinant strains of Salmonella minnesota and Escherichia coli expressing the genus-specific epitope of Chlamydia lipopolysaccharide. Eur J Biochem 222: 183–194, 1994.[ISI][Medline]
  18. Hori Y, Takeyama Y, Ueda T, Nishikawa J, Yamamoto M, and Saitoh Y. Impaired transport of lipopolysaccharide across the hepatocytes in rats with cerulein-induced experimental pancreatitis. Pancreas 16: 148–153, 1998.[ISI][Medline]
  19. Hsu DZ, Chiang PJ, Chien SP, Huang BM, and Liu MY. Parenteral sesame oil attenuates oxidative stress after endotoxin intoxication in rats. Toxicology 196: 147–153, 2004.[CrossRef][ISI][Medline]
  20. Jia HP, Kline JN, Penisten A, Apicella MA, Gioannini TL, Weiss J, and McCray PB Jr. Endotoxin responsiveness of human airway epithelia is limited by low expression of MD-2. Am J Physiol Lung Cell Mol Physiol 287: L428–L437, 2004.[Abstract/Free Full Text]
  21. Josza L, Lehto MU, Jarvinen M, Kvist M, Reffy A, and Kannus P. A comparative study of methods for demonstration and quantification of capillaries in skeletal muscle. Acta Histochem 94: 89–96, 1993.[ISI][Medline]
  22. Kaser A, Ludwiczek O, Waldenberger P, Jaschke W, Vogel W, and Tilg H. Endotoxin and its binding proteins in chronic liver disease: the effect of transjugular intrahepatic portosystemic shunting. Liver 22: 380–387, 2002.[CrossRef][ISI][Medline]
  23. Koutelidakis I, Papaziogas B, Giamarellos-Bourboulis EJ, Makris J, Pavlidis T, Giamarellou H, and Papaziogas T. Systemic endotoxaemia following obstructive jaundice: the role of lactulose. J Surg Res 113: 243–247, 2003.[CrossRef][ISI][Medline]
  24. Koyama I, Matsunaga T, Harada T, Hokari S, and Komoda T. Alkaline phosphatases reduce toxicity of lipopolysaccharides in vivo and in vitro through dephosphorylation. Clin Biochem 35: 455–461, 2002.[CrossRef][ISI][Medline]
  25. Marshall JC. Such stuff as dreams are made on: mediator-directed therapy in sepsis. Nat Rev Drug Discov 2: 391–405, 2003.[CrossRef][ISI][Medline]
  26. Meijer DK, Scholtens HB, and Hardonk MJ. The role of the liver in clearance of glycoproteins from the general circulation, with special reference to intestinal alkaline phosphatase. Pharm Weekbl Sci 4: 57–70, 1982.[ISI][Medline]
  27. Novikoff AB. Enzyme localizations with Wachstein-Meisel procedures: real or artifact. J Histochem Cytochem 15: 353–354, 1967.[ISI][Medline]
  28. Okada T, Zinchuk VS, and Seguchi H. Lipopolysaccharide administration increases acid and alkaline phosphatase reactivity in the cardiac muscle. Microsc Res Tech 58: 421–426, 2002.[CrossRef][ISI][Medline]
  29. Olinga P, Merema M, Hof IH, de Jong KP, Slooff MJ, Meijer DK, and Groothuis GM. Effect of human liver source on the functionality of isolated hepatocytes and liver slices. Drug Metab Dispos 26: 5–11, 1998.[Abstract/Free Full Text]
  30. Ortega-Cava CF, Ishihara S, Rumi MA, Kawashima K, Ishimura N, Kazumori H, Udagawa J, Kadowaki Y, and Kinoshita Y. Strategic compartmentalization of Toll-like receptor 4 in the mouse gut. J Immunol 170: 3977–3985, 2003.[Abstract/Free Full Text]
  31. Paik YH, Schwabe RF, Bataller R, Russo MP, Jobin C, and Brenner DA. Toll-like receptor 4 mediates inflammatory signaling by bacterial lipopolysaccharide in human hepatic stellate cells. Hepatology 37: 1043–1055, 2003.[CrossRef][ISI][Medline]
  32. Peterson AA and McGroarty EJ. High-molecular-weight components in lipopolysaccharides of Salmonella typhimurium, Salmonella minnesota, and Escherichia coli. J Bacteriol 162: 738–745, 1985.[Abstract/Free Full Text]
  33. Poelstra K, Bakker WW, Klok PA, Hardonk MJ, and Meijer DK. A physiologic function for alkaline phosphatase: endotoxin detoxification. Lab Invest 76: 319–327, 1997.[ISI]
  34. Poelstra K, Bakker WW, Klok PA, Kamps JA, Hardonk MJ, and Meijer DK. Dephosphorylation of endotoxin by alkaline phosphatase in vivo. Am J Pathol 151: 1163–1169, 1997.[Abstract]
  35. Pugin J, Stern-Voeffray S, Daubeuf B, Matthay MA, Elson G, and Dunn-Siegrist I. Soluble MD-2 activity in plasma from patients with severe sepsis and septic shock. Blood 104: 4071–4079, 2004.[Abstract/Free Full Text]
  36. Reddi AH and Anderson WA. Collagenous bone matrix-induced endochondral ossification hemopoiesis. J Cell Biol 69: 557–572, 1976.[Abstract/Free Full Text]
  37. Reichling JJ and Kaplan MM. Clinical use of serum enzymes in liver disease. Dig Dis Sci 33: 1601–1614, 1988.[CrossRef][ISI][Medline]
  38. Rensen PC, Oosten M, Bilt E, Eck M, Kuiper J, and Berkel TJ. Human recombinant apolipoprotein E redirects lipopolysaccharide from Kupffer cells to liver parenchymal cells in rats in vivo. J Clin Invest 99: 2438–2445, 1997.[ISI][Medline]
  39. Rietschel ET, Kirikae T, Schade FU, Mamat U, Schmidt G, Loppnow H, Ulmer AJ, Zahringer U, Seydel U, Di Padova and F. Bacterial endotoxin: molecular relationships of structure to activity and function. FASEB J 8: 217–225, 1994.[Abstract]
  40. Ruiter DJ, van der MJ, Brouwer A, Hummel MJ, Mauw BJ, van der Ploeg JC, and Wisse E. Uptake by liver cells of endotoxin following its intravenous injection. Lab Invest 45: 38–45, 1981.[ISI][Medline]
  41. Sheen-Chen SM, Chau P, and Harris HW. Obstructive jaundice alters Kupffer cell function independent of bacterial translocation. J Surg Res 80: 205–209, 1998.[CrossRef][ISI][Medline]
  42. Shimazu R, Akashi S, Ogata H, Nagai Y, Fukudome K, Miyake K, and Kimoto M. MD-2, a molecule that confers lipopolysaccharide responsiveness on Toll-like receptor 4. J Exp Med 189: 1777–1782, 1999.[Abstract/Free Full Text]
  43. Silverman MH and Ostro MJ. Bacterial Endotoxin in Human Disease: How Advances in Understanding the Role of Gram-Negative Bacteria and Endotoxin in Infectious Diseases and Complications May Improve the Development of Diagnostic and Treatment Options. Berkeley, CA: Xoma, 1998. [On-line: http://www.xoma.com:8080/pdf/kpmgendo.pdf].
  44. Sozinov AS. Systemic endotoxemia during chronic viral hepatitis. Bull Exp Biol Med 133: 153–155, 2002.[CrossRef][ISI][Medline]
  45. Su GL, Dorko K, Strom SC, Nussler AK, and Wang SC. CD14 expression and production by human hepatocytes. J Hepatol 31: 435–442, 1999.[CrossRef][ISI][Medline]
  46. Sugita H, Kaneki M, Tokunaga E, Sugita M, Koike C, Yasuhara S, Tompkins RG, and Martyn JA. Inducible nitric oxide synthase plays a role in LPS-induced hyperglycemia and insulin resistance. Am J Physiol Endocrinol Metab 282: E386–E394, 2002.[Abstract/Free Full Text]
  47. Takayama K, Qureshi N, Raetz CR, Ribi E, Peterson J, Cantrell JL, Pearson FC, Wiggins J, and Johnson AG. Influence of fine structure of lipid A on Limulus amebocyte lysate clotting and toxic activities. Infect Immun 45: 350–355, 1984.[Abstract/Free Full Text]
  48. Tillyer CR, Rakhorst S, and Colley CM. Multicomponent analysis for alkaline phosphatase isoenzyme determination by multiple linear regression. Clin Chem 40: 803–810, 1994.[Abstract/Free Full Text]
  49. Treon SP, Thomas P, and Broitman SA. Lipopolysaccharide (LPS) processing by Kupffer cells releases a modified LPS with increased hepatocyte binding and decreased tumor necrosis factor-alpha stimulatory capacity. Proc Soc Exp Biol Med 202: 153–158, 1993.[Abstract]
  50. Wiwanitkit V. High serum alkaline phosphatase levels, a study in 181 Thai adult hospitalized patients. [On-line: www.biomedcentral.com/content/pdf/1471-2296-2-2.pdf] 2001.




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