Division of Surgical Oncology, Massachusetts General Hospital and Harvard Medical School, Boston, Massachusetts 02114-2696
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ABSTRACT |
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Despite the central role of the liver in glutamine homeostasis in health and disease, little is known about the mechanism by which this amino acid is transported into sinusoidal endothelial cells, the second most abundant hepatic cell type. To address this issue, the transport of L-glutamine was functionally characterized in hepatic endothelial cells isolated from male rats. On the basis of functional analyses, including kinetics, cation substitution, and amino acid inhibition, it was determined that a Na+-dependent carrier distinct from system N in parenchymal cells, with properties of system ASC or B0, mediated the majority of glutamine transport in hepatic endothelial cells. These results were supported by Northern blot analyses that showed expression of the ATB0 transporter gene in endothelial but not parenchymal cells. Concurrently, it was determined that, whereas both cell types express glutamine synthetase, hepatic endothelial cells express the kidney-type glutaminase isozyme in contrast to the liver-type isozyme in parenchymal cells. This represents the first report of ATB0 and kidney-type glutaminase isozyme expression in the liver, observations that have implications for roles of specific cell types in hepatic glutamine homeostasis in health and disease.
amino acid transport; ATB0; system ASC; system B0; system N
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INTRODUCTION |
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THE PLEIOTROPIC ROLE OF circulating glutamine as an important metabolic fuel for dividing cells and as the major nontoxic shuttle of ammonia between tissues is underscored by its rapid cellular turnover rates (13) and presence at 0.6 mM in the plasma. It is well established that the liver plays a unique role in glutamine metabolism and that its ability to rapidly switch from net production to consumption of this amino acid is afforded by the differential position-dependent expression of specific glutamine-metabolizing enzymes in parenchymal cells (PCs) along the liver acinus (20). Portal and arterial glutamine is taken up by the large population (~95% of the total PCs) of "periportal hepatocytes" surrounding the proximal end of each sinusoid. Transported glutamine is subsequently hydrolyzed to ammonia and glutamate by the mitochondrial liver glutaminase isozyme, and the ammonia generated is detoxified by the urea cycle (44). In contrast, ammonia that escapes detoxification by the urea cycle is "scavenged" and utilized to generate glutamine via the enzyme glutamine synthetase (GS) by a small population (representing 5-7% of the total PCs) of "perivenous hepatocytes" surrounding the venous outflow. Thus the relative flux through each opposing pathway of this intercellular glutamine cycle (21) determines the net glutamine balance across the liver and may be controlled by the transport of glutamine across the plasma membrane (22, 32).
Although PCs constitute 70% of the total number of cells and nearly 90% of liver mass, sinusoidal nonparenchymal cells (NPCs) such as lipocytes and endothelial and Kupffer cells comprise ~30% of hepatic cellularity and 27% of its total plasma membrane content (3). Among the NPC types, sinusoidal endothelial cells that form the "sieve plates" are the most abundant, representing 2.5% of the lobular parenchyma (3) and from 38% (18) to 75% (31) of the total number of NPCs, based on protease-digested rat liver cell suspensions. Glutamine transport has been well characterized in isolated PCs from both rat (29) and human (5) livers. In both species, it is mediated by a unique Na+-dependent transporter that plays a central role in support of hepatic nitrogen metabolism (7, 22, 32) and has been termed system N for its narrow substrate specificity of only glutamine, histidine, and asparagine, amino acids with nitrogen-containing side chains. To date, transport processes in hepatic endothelial cells have focused exclusively on the receptor-mediated uptake of ligands such as lipoproteins, plasma proteins, and matrix components (8, 15), but, to our knowledge, no studies have yet examined amino acid transport processes in this second most abundant liver cell type. Given the important role of glutamine in hepatic physiology and metabolism, the studies presented here were undertaken to compare the uptake of this amino acid in liver endothelial cells (LECs) with the well-characterized PCs. The results show that the expression of isozymes for the transport and metabolism of glutamine is cell specific, reflecting the differential metabolic role of this amino acid in each cell type.
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MATERIALS AND METHODS |
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Radiolabeled
L-[3H]glutamine
and [-32P]dCTP were
obtained from DuPont NEN (Boston, MA). The digestive enzymes pronase
and collagenase were from Boehringer Mannheim (Indianapolis, IN), and
chemicals, perfusion media [suspension culture minimal essential
medium (S-MEM)], and unlabeled amino acids were from
Sigma (St. Louis, MO). Tissue culture medium and all additives were
from GIBCO BRL Life Technologies (Gaithersburg, MD), and supplies and
chemicals for scintillation spectrophotometry were from Packard
Instrument (Meriden, CT).
LEC isolation. Male Sprague-Dawley rats (500-700 g) were obtained from Charles River Laboratories (Wilmington, MA). Animals were housed in the Massachusetts General Hospital animal facility under controlled conditions of 12:12-h light-dark cycles and ad libitum access to chow and water. All experimental procedures were approved by the Massachusetts General Hospital Institutional Animal Care and Use Committee/Subcommittee on Research Animal Care, according to the guidelines in the Guide for the Care and Use of Laboratory Animals. To minimize nutritional influences on experimental results, all animals were subjected to an overnight fast before surgery. LECs were isolated by a modified pronase-collagenase digestion procedure established by others (18, 26, 30, 31), followed by density gradient centrifugation (18) and a final centrifugal elutriation step (18, 26, 30, 31). Briefly, rats were anesthetized intraperitoneally (75 mg/kg ketamine, 5 mg/kg xylazine, and 1 mg/kg acepromazine; Henry Schein, Port Washington, NY), laparatomy was performed, and the portal vein and inferior vena cava were cannulated with 16- and 14-gauge Teflon angiocatheters, respectively. The liver was cleared of blood by antegrade perfusion (20 ml/min) with 100 ml of warm (37°C) Ca2+-free S-MEM. Thereafter, the flow rate was reduced to 10 ml/min and the liver was first digested with pronase (2 mg/ml) in 100 ml of a Ca2+-containing balanced salt solution [BSS; containing (in mM) 140 NaCl, 10 HEPES (pH 7.4), 5 KCl, 5 glucose, 25 NaHCO3, and 0.5 CaCl2], followed by collagenase digestion (0.35 mg/ml) in 100 ml of Ca2+-containing S-MEM. The digested liver was removed, and cells were released by gentle agitation into 100 ml of BSS containing 30 mg pronase in a siliconized Erlenmeyer flask and then placed on a heat-calibrated (37°C) stir plate with a magnetic stir bar for an additional 30 min. In a subset of experiments, the pronase steps were eliminated and only collagenase was used for liver digestion; isolation of PC was performed in these preparations as described previously (17) for use in comparison studies. Afterward, 10 ml of fetal bovine serum were added to the mixture to help neutralize proteolytic activity, and the suspension was passed over eight-ply sterile gauze in a metal cell strainer into two 50-ml centrifuge tubes.
The suspension was subjected to two centrifugations at 50 g for 2 min to remove any remaining PC, and the supernatant containing the NPC was subjected to a final centrifugation at 500 g for 5 min. The NPC pellet was resuspended in 25 ml of S-MEM containing 10 µg/ml of DNase I, passed over a 40-µm cell strainer, and carefully layered on top of a discontinuous gradient of arabinogalactan density medium (Cellsep, Larex, St. Paul, MN) (18). After centrifugation at 2,500 g for 30 min, the NPCs were harvested from the 1.05- to 1.08-g/ml interface, diluted with S-MEM, and centrifuged at 500 g for 5 min. Hepatic lipocytes and cell debris were located at the medium to 1.05-g/ml interface. The NPC pellet was resuspended in 10 ml of S-MEM plus DNase I, passed over another 40-µm cell strainer, and loaded at a flow rate of 10 ml/min into the standard chamber of a Beckman JE-6B centrifugal elutriation rotor maintained at 2,500 rpm. The load eluate contained mostly cell debris and nonviable cells. The pump speed was increased to 25 ml/min, and the eluate was collected until the visible cell-to-medium interface in the chamber became clear. This fraction contained pure LECs, recognized by size, morphology in culture, and lack of peroxidase staining. The pump speed was increased to 40 ml/min, and the eluate was collected, which contained mostly LECs but some Kupffer cells as well. The "washout" fraction was also collected as the rotor was turned off and the pump speed increased to 60 ml/min. This fraction contained LECs as well but also Kupffer cells and aggregated cells. These results are consistent with those from other studies that utilized centrifugal elutriation to isolate LECs (26, 30, 31). The 25-ml/min cell fraction was utilized in all subsequent studies. To confirm that the 25-ml/min elutriated fraction was highly enriched in LECs, we injected a set of rats (50 µg/animal) with 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine-labeled acetylated low-density lipoprotein (Di-I-AcLDL; Molecular Probes, Eugene, OR) directly into the inferior vena cava 5-10 min before LEC isolation. Because LECs express a scavenger receptor for modified lipoproteins, this method has been utilized in the past to distinguish these cells from other liver cell types (34, 36). Once in culture, isolated cells were examined for Di-I-AcLDL content by means of an inverted phase-contrast microscope outfitted with an epifluorescence unit and rhodamine filter. Positive cells were identified by the presence of cytoplasmic red fluorescence, whereas uninjected rats were used as controls. The presence of contaminating Kupffer cells was assayed by morphology and a diaminobenzidine peroxidase staining method (4, 45).LEC culture. Once isolated, the LEC fraction was quantified and assessed for viability (typically >98%) by trypan blue exclusion on a hemacytometer and diluted in endothelial cell plating medium (GIBCO BRL) at a density of 1.2 × 106 cells/ml. The cells (0.5 ml/well) were placed in 24-well culture plates (Costar, Cambridge, MA) previously coated with type I rat tail collagen (Collaborative Biomedical Products, Bedford, MA) and allowed to attach for 2-3 h in a humidified atmosphere of 5% CO2-95% air at 37°C. Thereafter, the medium was changed to serum-free endothelial cell medium (GIBCO BRL), and the cells were maintained in monolayer culture until the transport assays were performed, typically 2-3 h later.
Amino acid transport assay.
Measurement of initial rate of glutamine uptake was carried out via the
cluster-tray method (19), as reported previously (5). The uptake of
L-[3,4-3H(N)]glutamine
was measured in the presence of specific amounts of unlabeled
L-glutamine as indicated. For
kinetic studies, the amount of unlabeled glutamine in the transport
buffer varied from 10 µM to 8 mM. After an initial two rinses with
Na+-free Krebs-Ringer phosphate
buffer (KRP), all transport measurements were carried out at 37°C
in either Na+-free KRP or
Na+-containing KRP and were
terminated after 1 min. Intracellular radiolabeled glutamine was
extracted with 0.2 ml/well of 0.2% SDS + 0.2 N NaOH; after 1 h, 0.1 ml
of the lysate was neutralized with 10 µl of 2 N HCl and subjected to
scintillation spectrophotometry (TopCount, Packard Instruments). The
remaining lysate was utilized for the determination of cellular protein
by the bicinchoninic acid method (Pierce Chemical, Rockford, IL). Rates
of glutamine transport were calculated from the counts per minute (cpm)
per sample and the specific activity of the uptake mix (in cpm/nmol) and normalized to cellular protein content in a Microsoft Excel spreadsheet program. Transport values obtained in the absence of
extracellular Na+ (diffusion and
Na+-independent uptake) were
subtracted from those in the presence of
Na+ (total uptake) to yield
Na+-dependent rates (reported in
units of nmol · mg
protein1 · min
1).
All transport values depicted are the average ± SD of four separate
determinations. Kinetic analysis of transport data was performed by
linear and nonlinear regression analysis in Cricket Graph (Computer
Associates, Islandia, NY).
Statistical analysis. Differences in specific measured parameters between experimental conditions were evaluated for statistical significance by paired t-test (Microsoft Excel) or ANOVA for multiple comparisons (Statview, Abacus Concepts, Berkeley, CA). Relative differences were considered significant at P < 0.050.
RNA isolation and Northern blot procedure.
Total cellular RNA was isolated from isolated LECs, PCs, or frozen
tissues (liver or kidney) by the one-step acid-phenol guanidinium procedure (11) using Trisolve reagent (Biotecx, Houston, TX), followed
by an additional acid-phenol, phenol-chloroform-isoamyl alcohol,
chloroform extraction and ethanol precipitation in the presence of
sodium acetate. Equal amounts of total RNA (20 µg), as determined
both spectrophotometrically and through ethidium bromide staining, were
fractionated by electrophoresis through denaturing 1% agarose gels
containing 0.2 M formaldehyde, transferred to nylon membranes by
capillary action, and ultraviolet cross-linked to the membrane. The
cDNAs utilized in this study to generate radiolabeled probes were
full-length rat ATB0
(rATB0;
Sal
I/Not I, 2.7-kb fragment) in pSPORT1
and kindly provided by Dr. Vadivel Ganapathy (27). Rat liver
glutaminase (GAL;
EcoR I, 0.6- and 1.4-kb fragments) in
pBluescript II SK was kindly provided by Dr. Malcolm Watford (39), rat
kidney type glutaminase (GAK;
Cla
I/Acc I, 1.3-kb fragment) in
pBluescript() was kindly
provided by Dr. Norman Curthoys (38), and rat GS (BamH I, 0.8-kb fragment), originally
derived from pGSRK-1 (9) in pBluescript II KS, was kindly provided by
Dr. John F. Mill. The cDNA inserts containing primarily coding sequence
were excised from the plasmid with appropriate restriction enzymes,
separated on agarose gels, excised, eluted, and used as templates to
generate [
-32P]dCTP-labeled
probes using a random primer labeling kit (Megaprime, Amersham,
Arlington Heights, IL) according to the manufacturer's protocol.
Hybridization with radiolabeled probe was performed overnight at
65°C in 5× sodium chloride-sodium phosphate-EDTA (SSPE) with 7.5× Denhardt's reagent, 0.5% SDS, and
0.1 mg/ml sheared herring sperm DNA, after the membrane was preblocked
for 2 h under the same conditions. Blots were washed at 55°C three
times each for 10 min in decreasing concentrations of SSPE and
increasing concentrations of SDS until 0.1× SSPE and 1.0% SDS
was reached. Autoradiograhic detection of the hybridization was
achieved by exposure of Fuji Medical X-ray film at
80°C. The
hybridized probe was stripped off the membrane by boiling in 0.1% SDS,
and the blots were reutilized for the Northern analyses of other genes.
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RESULTS |
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Purity of endothelial cells.
On attachment, cells eluted from the rotor at 25 ml/min displayed the
characteristic "spider web" cytoplasmic fenestrations characteristic of LECs, as well as cytoplasmic (red) fluorescence in
preparations from the animals injected with Di-I-AcLDL before liver
digestion (Fig. 1). In contrast, isolated
PCs failed to display this fluorescence as did uninjected animals (not
shown). Furthermore, the cells in the 25-ml/min elutriated fraction
failed to display the perinuclear peroxidase-dependent diaminobenzidine staining characteristic of Kupffer cells (4, 45). This staining was
evident, however, in (presumably) Kupffer cells in the 40-ml/min eluted
fraction plated on uncoated tissue culture dishes. Collectively, the
data suggest that the isolated NPC fraction utilized in these studies
is highly enriched in LECs. The cell purity and transporter characteristics (listed below) remained unaffected whether a
combination of pronase and collagenase or collagenase alone was
utilized.
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Glutamine transport in LECs.
Glutamine transport in isolated LECs was determined in the presence of
choline, Li+, or
Na+ as the major cation for 1 min.
The 1-min assay time was determined to be appropriate for initial rate
transport measurements, based on preliminary time courses as previously
described (5). As shown in Fig. 2, similar
to PCs the majority of glutamine uptake in LECs is
Na+ dependent (62%), whereas
Na+-independent and nonsaturable
uptake accounted for 24% and 14% of total transport values,
respectively. When the glutamine concentration was reduced to 10 µM,
the Na+-dependent component
increased to 78% of total transport values (0.090 ± 26 and 0.116 ± 19 nmol · mg
protein1 · min
1,
respectively), whereas at 600 µM glutamine (physiological levels) the
Na+-dependent contribution (2.26 ± 0.9 nmol · mg
protein
1 · min
1)
remained at 62% of the total uptake (3.62 ± 0.68 nmol · mg
protein
1 · min
1)
values. Figure 2 also shows that the LECs were intolerant to Li+-for-Na+
substitution to drive the concentrative uptake of 50 µM
L-glutamine, in contrast to PCs,
where such tolerance is a well-established system N functional
characteristic (5, 7, 29). These data suggest that the
Na+-dependent carrier expressed in
LECs is distinct from the system N glutamine transporter in PCs.
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LEC gene expression.
Recently, cDNAs were isolated that encode for system ASC (43) and
system B0 (27, 28) glutamine
transporters with characteristics similar to those reported here for
the LECs. The system ASC gene was isolated from a mouse cDNA library
and designated as ASCT2.
The system B0 genes were isolated
from human (27, 28) and rabbit (28) cDNA libraries, as well as from a
rat cDNA library (V. Ganapathy, unpublished observations),
and designated as ATB0 for
amino acid transporter B0. Both
the ASCT2 and
ATB0 genes encode for
glutamine transporters with nearly identical substrate specificities
and kinetic characteristics. The deduced amino acid sequences between
mouse ASCT2 (553 amino acids) and human ATB0
(541 amino acids) are highly similar (85%), with a 79% amino acid
identity. Both are members of the rapidly emerging glutamate transporter superfamily (24), which includes ASCT1, a system ASC
isoform that, in contrast to ASCT2 and
ATB0, does
not transport glutamine or asparagine (1, 37). Because our studies
focused on rat liver cells, we utilized the rat
ATB0 cDNA
(rATB0) to test for its
expression in the LECs by Northern blot analysis (Fig.
5). A single band of ~2.8 kb was observed
in both LEC and kidney but was not evident in RNA from whole liver or
isolated PCs. The apparent single 2.8-kb
ATB0 mRNA species and the lack of
detectable expression in liver (probably attributable to the
"swamping out" of LEC-derived by PC-derived RNA) and evident
expression in the kidney are consistent with results reported
previously for this glutamine transporter (27, 28).
Nonetheless, the expression of
ATB0 in the LEC is consistent with
the functional glutamine transport characteristics presented above;
unfortunately, the gene responsible for system N activity has not yet
been isolated.
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DISCUSSION |
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This study represents the first report on differences between liver cell types with respect to glutamine transport and the first report on the expression of ATB0 and GAK in normal liver, a finding previously masked by the preponderance of hepatic PC-derived RNA. Prior studies showed that LECs utilize glutamine as a respiratory substrate more readily than PCs (41). We show here that the concurrent expression of ATB0 and GAK in LECs probably contributes to this phenotype. The ammonia generated from the action of GAL is readily channeled into the urea cycle via carbamoyl phosphate synthetase-1, whereas the ammonia generated from GAK often escapes as free ammonia (44). Thus each glutaminase isozyme serves a different role in the metabolism of individual liver cell types. With respect to glutamine uptake in endothelial cells, this process has also been attributed to system "ASC-like" transporters in porcine pulmonary artery (25), rat lung microvascular (35), and human umbilical vein (10) endothelial cells. However, despite their common mesodermal origin, there are profound morphological and biochemical differences between LECs and vascular endothelium, including the pronounced fenestrae, which account for the "sieve-plate" morphology of the hepatic sinusoids (26). As demonstrated here, the system ASC-like glutamine uptake in LECs is attributable to the expression of the ATB0 gene (Fig. 5), which raises the possibility that this gene product may also mediate vascular endothelial cell glutamine transport previously described as system ASC by us (25, 35) and others (10).
System N-mediated glutamine across the plasma membrane of hepatocytes has been shown to represent a rate-limiting step in glutaminase-dependent glutamine metabolism, especially when intracellular utilization rates are accelerated (22, 23, 32). Similarly, it has been shown that system ASC-mediated glutamine transport plays an important role in metabolic regulation. For example, transport rates account for the 90% of glutamine turnover rates in cultured fibroblasts (14) and also govern growth rates in a human hepatoma cell line (6). Because Na+-dependent transporters such as systems N and ASC/B0 utilize the energy present in the Na+ electrochemical gradient to drive the concentrative uptake of substrates against their transmembrane gradient, their operation maintains cytoplasmic amino acid levels far above equilibrium. One reason that the Na+-dependent components of glutamine uptake play a quantitatively important role in governing cellular metabolic rates is that their activity is much higher than the Na+-independent components that allow net glutamine efflux (see Fig. 2). It is also the hepatic Na+-dependent components of glutamine uptake that increase during catabolic states, whereas there is little alteration in the Na+-independent components that allow dissipation of transmembrane amino acid gradients (33). In a well-designed study, Low and colleagues (32) quantitatively evaluated the contribution of system N to hepatocyte glutamine metabolism. The contributions of Na+-dependent and -independent transporters, mitochondrial transporter, glutaminase, proteolysis, and minor metabolic pathways were assessed for their role in observed glutamine utilization rates. Because GS is present in only 5% of the total liver PCs, this pathway was not considered in that study, although the capacity of this small population of cells to synthesize glutamine is enormous (Fig. 5 and Ref. 21). The authors concluded that, in the presence of physiological extracellular levels of histidine, system N possesses a significant flux control coefficient (0.51) and regulates glutaminase rates by adjusting cytoplasmic glutamine levels through electrochemical-dependent changes in its activity. System N activity was also found to be less than that of the mitochondrial transporter that supplies glutaminase, the major glutamine metabolizing enzyme in liver; in this sense, it can be considered a rate-limiting step in the regulation of glutamine catabolism. Together, it is apparent that cellular glutamine economy is governed by relative rates of synthesis and utilization, but Na+-dependent transporters contribute significantly to flux through pathways such as glutaminase.
The upregulation of hepatic transport activities during catabolic states may be especially crucial, when intracellular amino acid utilization rates are accelerated. Previous studies from our laboratory have demonstrated a marked increase in system N activity in PCs isolated from animals treated with bacterial lipopolysaccharide (LPS) (17), a response that probably contributes to the increase in net hepatic glutamine uptake observed during endotoxemia (2). Spolarics and Wu (40, 42) have studied LPS-induced effects on LEC pathways of reactive oxygen species detoxification, which are activated by this inflammatory agent and largely dependent on the pentose phosphate pathway and maintenance of intracellular glutathione levels. Because glutamine plays an essential role in glutathione metabolism, its uptake in LECs might be expected to be accelerated in response to LPS, similar to system N in PCs. Enhancement of ATB0 expression and activity in LECs during catabolic states may be necessary, given its much broader substrate specificity than system N, where glutamine must compete for uptake with more circulating amino acids. As shown in Fig. 5, however, LECs not only possess the ability to transport and metabolize glutamine but also have an appreciable capacity to produce it via GS. On the basis of the markedly different enzymes for glutamine transport and hydrolysis in LECs, the response of the LECs may deviate from that of the metabolically compartmentalized PCs during catabolic states. Nonetheless, the studies presented here should serve as the basis for future investigations into the contribution of LECs to global hepatic glutamine economy.
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ACKNOWLEDGEMENTS |
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We thank Dr. Vadivel Ganapathy, Dr. Malcolm Watford, and Dr. Norman Curthoys for providing the cDNAs used in this work. We also thank Dr. Nancy Reuter for contributions to some of the transport assays as well as Dr. Steve Abcouwer for assistance with the Northern blotting procedures.
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FOOTNOTES |
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This work was supported in part by a grant from the Harvard Clinical Nutrition Research Center (National Institute of Diabetes and Digestive and Kidney Diseases) Grant 1-P30 DK-40561 (to B. P. Bode) and a grant from the Deutsche Forschungsgemeinschaft, Bonn, Germany (Lo 599/1-1, to R. Lohmann).
Present address of R. Lohmann: Charité, Campus Virchow Clinic, Dept. of Surgery, Augustenburger Platz 1, 13353 Berlin, Germany.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: B. P. Bode, Massachusetts General Hospital, Surgical Oncology Research, Gray/Jackson 918, 55 Fruit St., Boston, MA 02114-2696 (E-mail: bode.barrie{at}mgh.harvard.edu).
Received 16 June 1998; accepted in final form 3 December 1998.
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