* Center for Neurosciences and Cell Biology of Coimbra, Department of Zoology, University of Coimbra, 3004517 Coimbra, Portugal; Department of Anatomy and Cell Biology, University of Minnesota School of Medicine, 1035 University Drive, Duluth, Minnesota 55812; and
Department of Biochemistry and Molecular Biology, University of Minnesota School of Medicine, 1035 University Drive, Duluth, Minnesota 55812
Received November 26, 2003; accepted January 20, 2004
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ABSTRACT |
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Key Words: chenodeoxycholic acid; ursodeoxycholic acid; combination; apoptosis; necrosis; mitochondria.
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INTRODUCTION |
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Previously, we have demonstrated that CDCA is toxic to hepatocytes by mechanisms involving mitochondrial dysfunction (Rolo et al., 2003a). Furthermore, we demonstrated that ursodeoxycholic acid (UDCA) potentiates CDCA-induced necrosis in primary cultured hepatocytes (Rolo et al., 2003b
), by inducing the mitochondrial permeability transition (MPT). This adds to the growing controversy regarding the utility of UDCA in treating cholestatic diseases (Gores et al., 1998
; Krahenbuhl et al., 1994
; Rodrigues et al., 1998
).
In recent years, considerable research regarding the subject of cholestasis has focused on the mechanisms by which hydrophobic bile acids induce apoptosis, and the putative protective effect of UDCA. It has been suggested that bile acids induce cell death via ligand-dependent death receptor pathways, especially those involving the Fas receptor (Higuchi et al., 2001; Miyoshi et al., 1999
). Other studies have implicated oxidative stress and mitochondrial dysfunction in bile acidinduced apoptosis (Graf et al., 2002
; Rodrigues et al., 1998
), specifically through induction of the MPT. Interestingly, hydrophobic and potentially toxic bile acids have been found to also activate survival pathways promoting cell proliferation (Qiao et al., 2002
; Rust et al., 2000
). The same survival pathway (activation of a mitogen-activated protein kinase, MAPK) has been implicated in protecting against UDCA toxicity, since UDCA has been shown to partially activate a pro-apoptotic cascade (Qiao et al., 2002
). It has been previously observed that UDCA induces apoptosis in hepatocellular carcinoma cells and potentiates photodynamic therapy (Kessel et al., 2000
; Oyama et al., 2002
).
The sum of these observations point to the necessity of systematic studies to fully understand the mechanisms of bile acids toxicity and the role of UDCA, either in preventing or potentiating bile acids toxicity. Toward this goal, we investigated whether CDCA-induced apoptosis is prevented by UDCA or its taurine-derivative, tauroursodeoxycholic acid (TUDC), and characterized the involvement of mitochondria in the process. Due to the interconnection between cell proliferation and programmed cell death, we also investigated the effects of bile acids on cell cycle.
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MATERIALS AND METHODS |
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Purified mouse anti-BrdU and purified mouse anti-ß-tubulin were purchased from Developmental Studies Hybridoma Bank (Iowa City, IA). Goat-anti-mouse Texas red was purchased from Jackson ImmunoResearch (West Grove, PA). Purified mouse anti-cytochrome c (clone 7H8.2C12) and purified mouse anti-human poly (ADP-ribose) polymerase (PARP, clone 4C10-5) were from PharMingen International (San Diego, CA). Goat anti-rabbit IgG was from Jackson ImmunoResearch Laboratories (West Grove, PA).
All other chemicals were of analytical grade and obtained from standard commercial sources.
Cell culture.
HepG2 cells were cultured in Minimum Essential Medium (MEM) Eagle (with 2 mM L-glutamine and Earle's BSS adjusted to contain 1.5 g/l sodium bicarbonate, 0.1 mM nonessential amino acids, and 1 mM sodium pyruvate), containing 10% fetal bovine serum. Cells were maintained in a humidified CO2 incubator at 37°C and passaged and harvested for experiments by detachement with 0.05% trypsin and 0.5 mM EDTA in phosphate-buffered saline (PBS); 8090% confluent cells were used for bile acid treatment. Control cultures received an equivalent amount of vehicle (water or ethanol).
Cell proliferation assay.
Cell proliferation was measured with sulforhodamine B as previously described (Skehan et al., 1990). Briefly, cells were seeded in 24-well plates at 5 x 104 cells/ml and allowed to attach and recover for 1 day prior to drug treatment. After addition of bile acids at the concentrations indicated in the legends to the figures, cells were cultured for up to 3 days without media change or drug replenishment. At various time points, media was removed and the wells rinsed twice with cold PBS. Adherent cells were fixed with cold 10% trichloracetic acid on ice for 30 min, washed, and stained with 0.5 % sulforhodamine B. Absorbance of the solubilized dye was measured at 540 nm in a Shimadzu UV1201 spectrophotometer.
Viability assay.
Cell viability was determined by the LIVE/DEAD Viability/Cytotoxicity Assay Kit (Papadopoulos et al., 1994) by flow cytometry. At 24 or 48 h following exposure to 50 µM CDCA and/or 50 µM UDCA or TUDC, cells were harvested by trypsinization and washed twice with PBS. Cells were resuspended in the working solution of calcein-AM (2 µM) and ethidium homodimer-1 (Eth-D1; 4 µM) and incubated for 30 min at 37°C. The cells were then analyzed with a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA) and CELLQuest software. Approximately 105 cells were analyzed for each time point and treatment.
Nuclear apoptosis assay.
HepG2 cell apoptosis was quantitated by determining the percentage of cells with nuclear morphologic changes of apoptosis detected by fluorescence microscopy of Hoechst 33342-stained fixed cells, as previously described (Holy, 2002). Briefly, cells were seeded on glass coverslips in 6-well plates at 5 x 104 cells/ml. At 24 or 48 h of treatment (50 µM CDCA and/or 50 µM UDCA or TUDC), coverslips were fixed and stored in formaldehyde and stained with 5 mg/ml Hoechst 33342. Coverslips were inspected and photographed on a Nikon Optiphot epifluorescence microscope using a Dage MTI SIT-68 video camera (Michigan City, IN). Two hundred and fifty cells from several randomly chosen fields were counted, and the number of apoptotic cells was expressed as a percentage of the total number of cells counted.
Immunocytochemistry.
Cells were seeded on glass coverslips in 6-well plates at 5 x 104 cells/ml. At 2 or 24 h of treatment (50 µM CDCA and/or 50 µM UDCA or TUDC), BrdU (5-bromo-2'-deoxyuridine) labeling solution (final concentration 10 µM in culture medium) was added and incubated for 3 h at 37°C. BrdU, a thymidine analogue, is a specific S-phase marker (Gratzner, 1982). Cells were then fixed and stored in cold (10°C) absolute methanol. At 24 and 48 h of treatment (50 µM CDCA and/or 50 µM UDCA or TUDC), cells for labeling with antibodies to ß-tubulin were also fixed. The procedure was essentially as previously described (Pierce et al., 1999
). Coverslips used for comparison of immunolabeling intensities between control and treated groups were all processed together and photographed at the same exposure times under identical conditions.
Cell cycle analysis.
Cell cycle analysis based on DNA content was performed as follows. At 24 h, following exposure to experimental treatments (50 µM CDCA and/or 50 µM UDCA or TUDC), cells were harvested by trypsinization and washed twice with PBS. Cells were then suspended in PBS, and 100 µl of RNase (1 mg/ml) and 100 µl of propidium iodide (100 mg/ml) were added to the cell suspensions and incubated at 37°C for 30 min. The cells were then analyzed with a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA) and ModFit cell cycle analysis software. Approximately 105 cells were analyzed for each time point and treatment.
Detection of cytochrome c release in cells.
To quantify cytochrome c release, immunoblot analysis of cytochrome c in the mitochondria and the cytosolic fractions were performed (Gottlieb and Granville, 2002). Briefly, at 48 h following exposure to experimental treatments (50 µM CDCA and/or 50 µM UDCA or TUDC), HepG2 cells were trypsinized, washed, and resuspended in 100 µl of ice-cold buffer (250 mM sucrose, 20 mM HEPES KOH (pH 7.5), 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM DTT, 1 mM PMSF, 1 µg/ml aprotinin, and 1 µg/ml leupeptin). After 30 min incubation on ice, cells were homogenized with a glass Dounce homogenizer. The protein concentration in the cytosolic and mitochondrial fractions, upon separation by centrifugation, was detected using the BCA protein assay kit. Equal amounts of protein were resuspended in SDSPAGE buffer and boiled in a water bath for 5 min. Lysates were stored at 80°C until further analysis. Proteins were separated under reducing conditions for 2 h at 120 V, as previously described (Pierce et al., 1999
).
Analysis of poly (ADP-ribose) polymerase cleavage.
Poly (ADP-ribose) polymerase (PARP) cleavage was detected by Western blot. Briefly, at the end of treatment (48 h of exposure to 50 µM CDCA and/or 50 µM UDCA or TUDC), cells were collected by trypsinization and washed with PBS. The final cell pellet was resuspended in an approximately equal volume of Tris-buffered saline (TBS). A threefold volume of SDS sample buffer (6% sodium dodecyl sulfate, 5% b-mercaptoethanol, 0.01% bromophenol blue in 40% glycerol) was added and the mixture boiled for 10 min. The boiled sample was passed through a 28-gauge syringe multiple times to shear the DNA and stored at 80°C until assayed, as previously described (Pierce et al., 1999).
Measurement of cellular caspase-9 activity.
Caspase-9 activity was quantified by measuring cleavage of a fluorometric peptide Ac-LEHD-AFC. Briefly, at the end of designated treatments (48 h of exposure to 50 µM CDCA and/or 50 µM UDCA or TUDC), cells were harvested and washed with PBS, and equal numbers of control or treated cells were lysed in cell lysis buffer (10 mM TrisHCl, pH 7.5, 10 mM NaH2PO4, 130 mM NaCl, 1% Triton X-100, and 10 mM NaPPi). The cell lysate was then incubated at 37°C with Ac-LEHD-AFC (10 µg/ml). After 2 h incubation, substrate cleavage leading to the release of the fluorescent AFC moiety (excitation 390 nm, emission 510 nm) was monitored by a Labsystems type 374 plate-reader. For blocking experiments, cell lysate was preincubated with caspase-9 specific inhibitor (Z-Leu-Glu/Ome-His-Asp(Ome)-FMK.TFA inhibitor) (10 µg/ml), prior to the addition of synthetic substrates. Changes in caspase activity were expressed as percent change from control, which was set as 100%.
Agarose electrophoresis for DNA fragmentation.
Cells (1 x 106) collected after treatments (48 h of exposure to 50 µM CDCA and/or 50 µM UDCA or TUDC) were washed in ice-cold PBS and resuspended in 450 ml digestion buffer (100 mM NaCl, 10 mM TrisHCl, pH 8, 25 mM EDTA, 0.5% SDS, 0.1 mg/ml proteinase K). RNase (final concentration 20 µg/ml) was then added, and incubation was continued at 37°C for 1 h. After incubation (50°C) overnight for complete digestion, DNA was extracted with an equal volume of phenol/chloroform/isoamyl alcohol and precipitated with ethanol. DNA fragmentation was detected by electrophoresis in 1.5% agarose gel containing 1 µg/ml ethidium bromide; 100-bp ladder molecular weight markers were used.
Measurement of mitochondrial membrane potential.
To monitor mitochondrial transmembrane potential (), HepG2 cells were loaded with 6.6 µM tetramethylrhodamine methyl ester (TMRM) in culture media, as previously described (Rolo et al., 2003b
). The different additions are shown in the legends to the figures.
Determination of ATP content.
The extraction of ATP from cells was performed as previously described (Ryll and Wagner, 1991), after 48 h incubation of HepG2 cells in culture with the compounds (50 µM CDCA and/or 50 µM UDCA or TUDC). ATP was quantitated by HPLC as previously described (Stocchi et al., 1985
) and the concentration expressed per million viable cells as determined by Trypan blue exclusion.
Statistical analysis.
Solvent controls (water or ethanol) were included within each experimental determination, and the numerical data are expressed as a percentage of the respective control (mean ± SEM, for 34 independent experiments). Ethanol itself had no effect on any of the parameters measured. Statistical significance was determined by the one-way ANOVA Student-Newman-Keuls post-t-test for multiple comparisons. A p value < 0.05 was considered statistically significant.
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RESULTS |
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At 24 h, no membrane alterations were observed (data not shown). However, at 48-h exposure to CDCA, there was a decrease in calcein-AM and an enhancement of Eth-D1 fluorescence (Fig. 2), indicating an increase in plasma membrane permeability. This was not surprising since it is known that, at late stages of apoptosis, plasma membrane becomes permeable. UDCA or TUDC (50 µM ) alone had no effect (Fig. 2). However, UDCA potentiated the changes caused by CDCA, suggesting an extensive population of damaged cells (Fig. 2). TUDC did not alter the toxicity caused by CDCA (Fig. 2).
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We performed an additional study to evaluate DNA synthesis, measuring bromodeoxyuridine (BrdU) incorporation (Fig. 3B, lower panel). Less then 1% of cells treated with 50 µM CDCA for 24 h incorporate BrdU. The 24-h exposure was compared with an earlier short (2 h) exposure to establish a time-dependent effect. We conclude that CDCA at 24 h induced cell growth inhibition through induction of cell cycle arrest, because no significant apoptosis is evident in these cell cultures as shown in Figure 3B (upper panel) with cells labeled with Hoechst 33342.
Effects on Chromatin Condensation and Tubulin Organization
In the sequence of the results described, we evaluated DNA changes by Hoechst 33342 staining following cell exposure to the different compounds for 48 h. Since apoptotic cells undergo a marked change in morphology, we also assessed alterations in tubulin organization, a component of the cytoskeleton.
A study of the nuclear changes, by staining with Hoechst 33342 dye, demonstrated that 50 µM CDCA induced nuclear changes typical of apoptosis in 40% of the cells by 48 h (Figs. 4A and 4B). Disorganization of tubulin was also observed (Fig. 4A).
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Effects on DNA Fragmentation
DNA fragmentation was used as another criterion to distinguish between apoptosis and necrosis. Consistent with the previous nuclear changes, cells treated with 50 µM CDCA or 50 µM CDCA plus 50 µM TUDC for 48 h started to show DNA fragmentation (Fig. 5). In UDCA plus CDCAtreated cells, on the other hand there was no cleavage of DNA that could be detected by gel electrophoresis (Fig. 5). No DNA fragmentation was observed in cells treated with UDCA or TUDC alone. These results further support a prominent role for necrotic cell death upon exposure to the combination UDCA plus CDCA, although apoptotic nuclei were observed with Hoechst 33342 staining.
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DISCUSSION |
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Numerous studies have demonstrated that cells undergoing apoptosis exhibit a decrease in mitochondrial membrane potential, preceding nuclear signs of apoptosis (Ferri and Kroemer, 2001), suggesting that early alterations of mitochondrial function may be important for initiating the apoptotic process. Our results demonstrate the activation of the mitochondrial apoptotic pathway in a hepatoma cell line following CDCA exposure, involving MPT induction. Such an event was shown by the early depolarization of mitochondrial membrane potential that is inhibited by CyA. In fact, impairment of mitochondrial function has been previously demonstrated to be a key mechanism by which bile acids cause cell dysfunction (Krahenbuhl et al., 1994
; Rolo et al., 2000
, 2003b
). Cytochrome c release and caspase-9 activation, with subsequent PARP cleavage and nuclear condensation, followed the onset of MPT by CDCA. It is well known that activated caspase-9 triggers a cascade that culminates in the activation of caspase-3, which cleaves several substrates, resulting in chromosomal DNA fragmentation and cellular morphologic changes characteristic of apoptosis (Cai et al., 1998
). Activation of caspase-9 usually occurs downstream of cytochrome c release from mitochondria and may be preceded by mitochondrial translocation of Bax (Narita et al., 1998
). A previous study by Rodrigues and coworkers, reported that deoxycholic acidinduced apoptosis in primary rat hepatocytes was associated with decreased mitochondrial transmembrane potential and increased mitochondria-associated Bax protein (Rodrigues et al., 1999
). Indeed, Bax activation has been proposed as an integrator of diverse pro- and anti-apoptotic signals, cell fate being decided by the resultant release of cytochrome c (Brenner and Kroemer, 2000
). This would explain the role for the Fas pathway as a common mechanism by which cytotoxic bile acids also induce apoptosis (Sodeman et al., 2000
), which was not evaluated in the present work. Interestingly, a recent work by Schoemaker and coworkers (Schoemaker et al., 2003
) has shown that the glycine derivative of CDCA induces apoptosis in a mitochondria-controlled pathway in which caspase-8 is activated in a FADD-independent manner. Therefore, apoptotic pathways are differentially activated by the distinct bile acids.
From this study, UDCA potentiates MPT induction in HepG2 cells by CDCA, as previously shown in isolated mitochondria (Rolo et al., 2000). The ability of UDCA to enhance mitochondrial membrane depolarization, sensitive to CyA, was associated with a much greater loss of ATP. Apoptotic changes in CDCA-treated cells occurred without a massive disruption of the plasma membrane as indicated by calcein-AM retention, Eth-D1 exclusion, and ATP loss. Thus, it appears that apoptosis is activated at low CDCA concentration and occurs after a long time of exposure, when compared with the combined bile acid cytotoxicity. Interestingly, cells exposed to UDCA alone underwent only a slight decrease in mitochondrial membrane potential with no significant change in ATP content, suggesting that cytotoxicity induced by UDCA is only expressed when CDCA is present. Such an effect may occur by potentiation of MPT induction. One explanation for this difference could be that induction of the MPT in a small fraction of mitochondria could activate the apoptotic process, whereas induction of mitochondrial transition in a large portion of mitochondria could cause bioenergetic collapse, as shown by ATP depletion, beyond which apoptosis could not proceed, leading to necrosis. Previously, we have shown that CDCA plus UDCA induces a decrease in mitochondrial respiration and activates the mitochondrial permeability transition (Rolo et al., 2000
, 2003b
), resulting in a bioenergetic failure, with cell death showing necrotic characteristics (Rolo et al., 2003b
). These changes occurred over a rapid time course (4 h) and following exposure to higher concentrations of bile acids (Rolo et al., 2003b
).
It has been proposed that caspase inactivation in lethally injured cells switches the mode of cell death from apoptosis to necrosis (Lemaire et al., 1998). Although cytochrome c release in CDCA plus UDCAtreated cells was evident, no caspase activity was observed. Such results suggest that ATP depletion plays a key role in caspase inactivation and loss of plasma membrane integrity. It may also explain the inhibition of other typical features of apoptosis in cells exposed to CDCA plus UDCA.
It is clear that bile acids, although at higher concentrations than used in this work, stimulate the generation of reactive oxygen species in isolated rat hepatocytes, primary cultured hepatocytes, and purified hepatocyte mitochondria (Rodrigues et al., 1998; Sokol et al., 1995
). Yerushalmi and collaborators reported that, in glycochenodeoxycholic acidinduced apoptosis, oxidative stress was a key mediator of the mitochondrial permeability transition (Yerushalmi et al., 2001
). We do not exclude the oxidation of cysteine residues in caspases as the mechanism responsible for their inactivation.
The role of UDCA in hepatic apoptosis is controversial. On one hand, UDCA has been described as decreasing apoptosis induced by deoxycholic acid and tumor growth factor-ß in both hepatic and nonhepatic cells (Rodrigues et al., 1998, 1999
). The probable mechanism was suggested to be via modulation of mitochondrial function and membrane stability, although a role of alteration of the bile acid pool was not ruled out (Rodrigues et al., 1998
). Other authors have described potentiation of cell dysfunction induced by UDCA. UDCA induces apoptosis in hepatocellular carcinoma cells, mediated by increased levels of Bax in mitochondria and cytochrome c in the cytosol (Oyama et al., 2002
). Preincubation of murine leukemia and hepatoma cells with UDCA potentiated the effects of photodynamic therapy (Kessel et al., 2000
). Such action involved effects at the level of mitochondrial depolarization, release of cytochrome c into the cytosol, and activation of caspase-3. Glyco- and tauro-conjugates of UDCA displayed the same action (Kessel et al., 2000
). In contrast, TUDC proved to be effective in reducing both GCDC-induced apoptosis and cytolysis (Benz et al., 1998
). The condition that UDCA exerts such an effect, if present before but not upon irradiation (Kessel et al., 2000
), supports our suggestion that UDCA per se does not cause mitochondrial membrane damage but lowers the threshold for the action of other compounds. Another interesting finding was the observation that UDCA activates a pro-apoptotic cascade, which is generally inhibited by a simultaneous and stronger activation of the MAPK survival pathway (Qiao et al., 2002
). These discrepant results may also be dependent on the different characteristics of the cell culture and the different expression of regulatory proteins. For example, S-adenosylmethionine has been described as an anti-apoptotic agent in cultured rat hepatocytes but pro-apoptotic in human hepatoma cells (Ansorena et al., 2002
).
Interestingly, besides a key regulatory role in signal transduction pathways, anti-apoptotic properties attributed to TUDC have been associated with potent mitochondrial membrane-stabilizing effects, preventing the binding of Bax to the outer mitochondrial membrane (Rodrigues et al., 2001). However, previous studies with the goal to characterize the common apoptotic signaling pathways in human hepatoma cells have shown that p53 and c-Myc play a more important role in the apoptotic signaling pathway in HepG2 cells than the Bcl-2 gene family (Jiang et al., 1996
). Therefore, the lack of effect of TUDC in CDCA-induced apoptosis may be the result of activation of different apoptotic pathways. However, since the hepatoblastoma cell line HepG2 is unable to take up bile acids via sodium-dependent pathways (the main system for the uptake of conjugated bile acids) because Ntcp is not expressed (Boyer et al., 1993
), this may also be the explanation for TUDC behavior in this model. Sodium-independent hepatocellular uptake of bile salts is mediated by several members of the OATP family of membrane transporters, which are multispecific transporters that mediate hepatocellular uptake of a vast variety of other amphipathic organic compounds in HepG2 cells.
TUDC is the taurine conjugate of UDCA, and differential effects on the induction of apoptosis may be a consequence of the altered physical-chemical properties. For example, amidation is known to significantly reduce the cytotoxicity of the more lipophilic forms of unconjugated bile acids, both in hepatocytes (Scholmerich et al, 1984) and bile duct cells. Sarbu and coworkers (2001) published on the lipophilic character of bile acids and their tauro-conjugates and showed that the main difference between these compounds is their polarity and acidity. Tauro-conjugates are more polar and acidic and, thereby, substantially less lipophilic than unconjugated bile acids, which appear to be important determinants of cytotoxicity.
In summary, the results of this study demonstrate that activation of apoptosis by CDCA involves an early loss of mitochondrial membrane potential. Cytochrome c release and activation of the caspase cascade probably follow this initial event. Coincubation with UDCA appears to induce necrosis as the predominant form of cell death, by a mechanism that involves extensive mitochondrial dysfunction. In view of this, although CDCA, by itself and at low concentrations, is an inducer of apoptosis, UDCA potentiates CDCA cell toxicity not by increasing apoptosis, but by diverging to necrotic pathways.
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Department of Biochemistry and Molecular Biology, University of Minnesota School of Medicine, 1035 University Drive, Duluth, MN 55812-2487. Fax: (218) 726-8014. E-mail: kwallace{at}d.umn.edu
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