Department of Cardiovascular Biology, Aventis Pharmaceuticals Research and Development, Collegeville, Pennsylvania 19426-0994
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ABSTRACT |
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The proposed
mechanism for the triglyceride (TG) lowering by fibrate drugs is via
activation of the peroxisome proliferator-activated receptor-
(PPAR
). Here we show that a PPAR
agonist, ureido-fibrate-5 (UF-5), ~200-fold more potent than fenofibric acid, exerts
TG-lowering effects (37%) in fat-fed hamsters after 3 days at 30 mg/kg. In addition to lowering hepatic apolipoprotein C-III (apoC-III)
gene expression by ~60%, UF-5 induces hepatic mitochondrial
carnitine palmitoyltransferase I (CPT I) expression. A 3-wk rising-dose treatment results in a greater TG-lowering effect (70%) at 15 mg/kg
and a 2.3-fold elevation of muscle CPT I mRNA levels, as well as
effects on hepatic gene expression. UF-5 also stimulated mitochondrial
[3H]palmitate
-oxidation in vitro in human hepatic and
skeletal muscle cells 2.7- and 1.6-fold, respectively, in a
dose-related manner. These results suggest that, in addition to
previously described effects of fibrates on apoC-III expression and on
peroxisomal fatty acid (FA)
-oxidation, PPAR
agonists stimulate
mitochondrial FA
-oxidation in vivo in both liver and muscle. These
observations suggest an important mechanism for the biological effects
of PPAR
agonists.
fibrates; carnitine palmitoyltransferase I; nuclear receptors; gene expression; peroxisome proliferator-activated receptor
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INTRODUCTION |
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THE PEROXISOME
PROLIFERATOR-ACTIVATED RECEPTOR (PPAR) is so named
because of its initial identification as the molecular mediator of the
peroxisome proliferation response to a number of chemical stimuli,
including fibrates, in mice (15, 17). The regulation of
genes involved in peroxisomal fatty acid (FA) -oxidation by PPAR
activators is well recognized (23). However, the
peroxisomal response is likely to be rodent specific and may not occur
in humans (14, 31).
Elevated plasma triglyceride (TG) concentrations constitute an
independent risk factor for coronary artery disease (11, 13). Fibrate drugs and FAs are believed to be weak PPAR
agonists, and PPAR
is likely to mediate the hypolipidemic effects of
fibrate TG-lowering therapy. One demonstrated mechanism for the
TG-lowering effects of fibrates that is likely to occur in humans is
via reduction of hepatic apolipoprotein C-III (apoC-III) transcription
and synthesis (28). Because apoC-III is thought to inhibit
very low density lipoprotein (VLDL)-TG hydrolysis by lipoprotein lipase
and to inhibit uptake of VLDL remnants (1, 26, 32), a
reduction in apoC-III synthesis would be predicted to result in a
lowering of plasma TG concentrations.
In addition to apoC-III, molecular studies have implicated PPAR in
the regulation of a number of genes involved in mitochondrial FA
-oxidation; however, the PPAR responsiveness of such genes in vivo
is less well studied. Carnitine palmitoyltransferase I (CPT I)
catalyzes the transfer of FA from CoA to carnitine, allowing the
initial transport of fatty acids into mitochondria for
-oxidation. Its activity and expression are highly regulated and rate limiting. Eicosapentaenoic acid and fenofibrate administration increased mitochondrial CPT I and II activities in rabbits (9).
Shunting of FAs toward
-oxidation would be expected to result in
decreased substrate availability for TG synthesis in liver, presumably
resulting in a reduction of VLDL-TG secretion. Although fibrate drugs
and FAs are believed to exert their effects on gene regulation via PPAR
activation, TG lowering and regulation of mitochondrial FA
-oxidation genes have not been directly demonstrated with the use of
a potent, bona fide PPAR
agonist. We used a ureido-fibrate analog
(UF-5), shown to stimulate microsomal FA-
hydroxylation and to lower
VLDL cholesterol in rats (12), for this purpose.
The fat-fed hamster represents a potentially important model of nondiabetic hypertriglyceridemia. Unlike other rodents, hamsters respond to fat feeding with a greater than twofold increase in plasma TG levels (29). Lipoprotein metabolism in hamsters may more closely reflect that of humans than of rats or mice (29), but hamsters are not generally considered responsive to classical fibrates (16). Here we describe the effects of UF-5 on TG metabolism in vivo in a hamster model of hypertriglyceridemia and on FA catabolism in vitro.
PPAR-mediated responses have been traditionally studied in liver,
but human and rat skeletal muscle expresses high levels of PPAR
(21), and in humans, skeletal muscle may be the major site
of PPAR
expression. We therefore hypothesized that a potent PPAR
agonist would stimulate mitochondrial FA
-oxidation in muscle as
well as in liver.
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METHODS |
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Chemicals. 2-(4-(2-(N'-(4- fluorophenyl)-N-heptylureido)ethyl)phenoxy)-2-methylpropionic acid was synthesized as previously described (12). Fenofibrate was obtained by hydrolysis of fenofibrate methyl ester (Sigma, St. Louis, MO). Wy-14643 was from BioMol (Plymouth Meeting, PA). Etomoxir was kindly provided by Dr. H. P. O. Wolf (Allensbach, Germany).
In vivo protocol.
Male Golden Syrian hamsters (Harlan Sprague Dawley, Madison, WI),
weighing 120-135 g, were group housed with a 12:12-h light-dark cycle. Hamsters were placed on a high-fat high-cholesterol diet (0.05%
cholesterol, 10% coconut oil; Dyets, Bethlehem, PA) for ~2 wk before
treatment with UF-5 or vehicle and continued on this diet throughout
the treatment period. UF-5 was prepared by sonication with vehicle
(0.5% methylcellulose, 0.2% Tween 80) and administered twice daily by
gavage in a rising-dose fashion (7.5, 15, and 30 mg/kg,
n = 7 animals/group) with each dose given for a 1-wk
period, resulting in a total of 3 wk of treatment with incremental
doses. Vehicle was administered at 5 ml/kg. In a second study, vehicle, UF-5 (7.5, 15, and 30 mg/kg), and fenofibric acid (30, 60, and 120 mg/kg) were administered twice daily to fat-fed hamsters
(n = 6/group) with a similar 3-wk rising-dose protocol.
Blood samples, removed under CO2 narcosis, were obtained at
specified times throughout the studies and were analyzed for
triglycerides, cholesterol, and compound. All blood samples, except
where indicated, were removed from animals fasted for 17 h. All
samples were taken 45-60 min after dosing. Hamsters were
terminated by CO2 overdose, and select tissues (liver and
soleus muscle) were removed, blotted, weighed, flash-frozen in liquid
N2, and frozen (70°C) for subsequent RNA analyses. All
animal protocols were approved by the Institutional Animal Care and Use Committee.
Quantitation of plasma UF-5 concentrations. Plasma samples were extracted with 0.1 µg/ml internal standard solution in acetonitrile on Porvair filtration disks. One hundred microliters of 10 mM ammonium acetate pH 3.5 were added and samples injected onto liquid chromatography-mass spectrometry/mass spectrometry in ionspray (positive) mode with magnetic resonance monitoring. Chromatography was performed on Luna C8 (2) 30 × 4.6 mm × 3 µm (Phenomenex), with 15-90% acetonitrile-ammonium acetate gradient as mobile phase with flow rate of 1 ml/min. The difluoro analog (12) of UF-5 was used as internal standard.
Quantitation of plasma TG, cholesterol, and glucose concentrations. Plasma TG concentrations were measured by the peroxidase method using the Sigma Diagnostic assay kit according to manufacturer's instructions. Plasma high-density lipoprotein (HDL) cholesterol was determined enzymatically after precipitation of apoB-containing lipoproteins by the phosphotungstic acid-Mg2+ method with Autokit Cholesterol (Wako Pure Chemical Industries, Osaka, Japan) and spectrophotometry with an automatic analyzer (Hitachi model 7050) at 600 nm. Plasma low-density lipoprotein (LDL) cholesterol was measured with the LDL Direct assay kit (Wako Pure Chemical Industries) (cholesterol esterase/cholesterol oxidase method), and the resulting H2O2 was measured colorimetrically at 585 nm. Plasma glucose was quantitated by the glucose oxidase-peroxidase method using the Sigma Diagnostic assay kit according to the manufacturer's instructions.
Cell culture. HepG2 cells were obtained from the American Type Culture Collection (ATCC; Manassas, VA) and cultured in DMEM-10% FCS-1% penicillin-streptomycin. Human primary skeletal muscle cells were obtained from Clonetics (San Diego, CA) and cultured in SkBM, supplemented with SkGM Singlequots.
Northern blotting.
RNA was extracted from hamster soleus muscle or liver with Trizol
reagent according to the manufacturer's protocol. Total RNA was
subjected to Northern blotting onto Nytran membranes (Schleicher & Schuell, Keene, NH). A probe for rat apoC-III was cloned exactly as
previously described (28). Human liver and muscle CPT I
probes (accession nos. R28631 and W85710, respectively) were excised from pT7T3 by EcoR I/Not I and EcoR
I/Pac I digestion, respectively. A probe corresponding to
nucleotides 760-964 of the rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH) coding sequence was cloned by PCR with primers 5'-CATCAAGAAGGTGGTGAAGC-3 (forward) and 5'-ACCCTGTTGCTGTAGCCATA-3 (reverse) into PCR2.1 and excised with EcoR I. A probe to
the mouse homolog of human S10 (accession no. NM001014) was prepared by
PCR of mouse liver cDNA using primers corresponding to nucleotides 17-170. Full-length human PPAR was subcloned from pSG5-hPPAR
(a kind gift from Dr. Bart Staels) into pcDNA3.1. To create
pcDNA3.1-hPPAR
, Bluescript SK
, containing a full-length hPPAR
-1
(ATCC), was digested with Asp718, filled in, digested with Not
I, and then ligated into pcDNA3.1 digested with Not
I/Kpn I (filled in). Probes for hPPAR isoforms were
excised from pcDNA3.1. Probes were labeled with the Random Primers DNA
Labeling System (Life Technologies, Rockville, MD) and
[
32P]dCTP (Amersham, Buckinghamshire, England). Blots
were hybridized with probes (as indicated in Figs. 1-5) with
ExpressHyb (Clontech, Palo Alto, CA) and washed according to the
manufacturer's protocol. After exposure of membranes to X-ray film,
signals were quantitated by densitometry (Personal Densitometer SI,
Molecular Dynamic, Sunnyvale, CA). Blots were stripped for reprobing by
boiling 2 × 10 min in 0.5% SDS.
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Western blotting.
Cell proteins were extracted from 100-mm dishes with PBS-1% Triton
X-100-5 mg/ml NaEDTA-1 mM phenylmethylsulfonyl fluoride. Extracts
were sonicated and centrifuged at 20,000 g for 15 min at
4°C. Proteins were resolved by electrophoresis on 10% SDS-PAGE gels,
transferred onto nitrocellulose membranes, and probed with rabbit
polyclonal antibodies against human PPAR (6) or PPAR
(kind gift of Bart Staels) by use of the Western Breeze (Novex, San
Diego, CA) hybridization/detection system. pcDNA3.1-hPPAR
and -
positive controls were transcribed and translated in vitro with the
Promega (Madison, WI) system as per manufacturer's instructions. Protein was quantitated by the Bradford method with the Bio-Rad (Hercules, CA) reagent according to manufacturer's instructions, with
BSA as standard.
Cellular FA -oxidation assay.
The rate of cellular
-oxidation of
[9,10(n)-3H]palmitic acid (52 Ci/mmol, Amersham) was
measured as 3H2O release, as previously
described (20). For cell incubations, [3H]palmitic acid was used at a final concentration of 22 µM in Hanks' balanced salt solution-0.5% free FA-free BSA (Sigma)
by dilution with unlabeled palmitic acid (Sigma). Cell DNA content was
quantitated with pico Green double-strand DNA quantitation reagent
(Molecular Probes, Eugene, OR) according to the manufacturer's instructions. Linearity of palmitate
-oxidation with time and cell
number was established by plating increasing numbers of HepG2 cells and
measuring cellular FA
-oxidation after 48 h of growth. After 1- or 2-h incubation with [3H]palmitate, cell supernatants
were assayed for palmitate
-oxidation product. On the basis of these
data, cells were plated at 1.2 × 105 cells/well, and
a 2-h incubation with substrate was used for experiments. UF-5 was
prepared as 100 mM stock in DMSO and added once every 24 h. In
some experiments, 40 µM etomoxir (2) was included either
as a 24-h preincubation or during the latter 24 h to inhibit CPT I.
Statistics.
Data are presented as means ± SE. Means were compared with
Student's t-test. In Figs. 1-5 and Tables 1 and 2, a
P value of <0.05 was considered significant.
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RESULTS |
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Effects of UF-5 on FA and
TG metabolism in fat-fed hamsters.
Recent reports (4, 12) demonstrated that UF-5 is a potent
PPAR activator with an EC50 value of 400 and 30 nM for
human and mouse PPAR
, respectively, in cell-based transactivation assays.
Effects of treatment duration on UF-5-induced lipid metabolism changes. To see how the treatment duration affects TG lowering and gene expression, fat-fed hamsters were treated for 3 days with UF-5 or vehicle. TG lowering with 30 mg/kg treatment of the nonfasted hamster was 37% after 3 days, or ~50% of the effect of the rising-dose treatment (Table 1). However, suppression of hepatic apoC-III levels was similar to that achieved after the rising-dose treatment (56 vs. 63%; Fig. 2A), and stimulation of hepatic CPT I expression was even greater (160 vs. 37%; Fig. 2A). Muscle-type CPT I-to-GAPDH mRNA ratio was unaffected in hamsters treated for 3 days with 30 mg/kg UF-5 (Fig. 2B). To see whether any effect might have been underestimated due to upregulation of GAPDH by UF-5, blots were stripped and reprobed with a murine S10 corresponding to a ribosomal protein. Muscle CPT I/S10 mRNA levels were not significantly different among any of the treatment groups (Fig. 2B).
Effects of UF-5 compared with fenofibric acid on
lipid metabolism in hamsters.
Next we compared the effects of fenofibric acid (FF), a very weak
PPAR activator (4), to UF-5 in vivo. As expected,
administration of rising doses of UF-5 to fat-fed hamsters lowered
plasma TG concentrations by >50%, whereas FF at the doses tested did
not (Table 2). UF-5, but not FF
treatment, also lowered plasma LDL cholesterol (Table 2), consistent
with putative inhibitory effects on VLDL secretion. Hepatic apoC-III
mRNA levels in hamsters treated for 3 wk with rising doses of UF-5 or
FF were reduced by ~50% when corrected for GAPDH, compared with
vehicle-treated hamsters (Fig.
3A). Soleus muscle CPT I mRNA
levels were increased approximately twofold in UF-5-treated hamsters
but were not affected in FF-treated animals (Fig. 3B).
UF-5 stimulates mitochondrial FA-
oxidation in vitro.
These results suggested that, in addition to previously described
effects of fibrates on apoC-III expression and on peroxisomal FA
-oxidation, PPAR
agonists might also exert TG-lowering effects through stimulation of mitochondrial FA
-oxidation. To further explore the ability of PPAR
agonists to stimulate FA
-oxidation in vivo, we studied cell types representative of tissues carrying out
high rates of FA
-oxidation, namely HepG2 and hSKMC. To assess the
suitability of the cell types for these experiments, the presence of
PPAR
mRNA and protein was first established. mRNA for hPPAR
was
in greater abundance in human skeletal muscle tissue than in liver
(Fig. 4A). HepG2 and hSKMC
expressed PPAR
mRNA (Fig. 4A) and also contained
immunoreactive PPAR
protein (Fig. 4B) in hSKMC until 7 passages.
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DISCUSSION |
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It is widely believed that the lipid-lowering effects of fibrates
are mediated through PPAR (22). Fibrates have been
shown retrospective of their clinical efficacy to be very weak PPAR
agonists, with unmeasurable binding affinities to PPAR
and with EC50 values in cell-based reporter assays in the tens of
micromolar range (4). It has been hypothesized that more
potent, directed PPAR
agonists would exert more powerful TG-lowering
effects (27). The data reported herein support the
hypothesis with a potent bona fide PPAR
agonist, UF-5
(4), in a fat-fed hamster model of nondiabetic
hypertriglyceridemia. Greater than 50% plasma TG lowering with UF-5
was seen at doses an order of magnitude lower than those typically
necessary for clinically used fibrates. These effects are likely to be
mediated through PPAR
, given that selectivity for murine PPAR
isoform activation of UF-5 is 26-fold for PPAR
over PPAR
(4).
A further objective of this study was to explore the potential
mechanism for TG lowering of PPAR agonists. Consistent with the
effects of fenofibrate in rats (28), we observed a sharp reduction in hepatic apoC-III expression in fat-fed hamsters. In
addition to effects on hepatic apolipoprotein expression, we show here
that PPAR
agonist treatment influences CPT I expression. Two CPT I
isoforms, liver (3, 7) and muscle (33, 34), have been cloned and characterized. Previous molecular evidence has
implicated PPAR
in regulation of the muscle isoform. The promoter of
human muscle-type CPT I is stimulated by FAs, a response mediated by a
PPRE to which PPAR
binds (19, 35). Oleate increases CPT
I expression in cardiac myocytes (2). More limited
information also implicates PPAR in regulation of the liver isoform;
clofibrate and FAs increase CPT I mRNA expression in fetal rat
hepatocytes (5). In this study, UF-5 treatment of fat-fed
hamsters upregulated both hepatic and muscle CPT I.
PPAR is also implicated in regulation of other genes involved in
mitochondrial FA
-oxidation. The promoters of medium-chain acyl-CoA
dehydrogenase and rat mitochondrial HMG-CoA synthase are PPAR
responsive (10, 24). Expression of acyl-CoA synthase (18, 25) and fatty acid transfer protein (8,
18) is induced by fibrates in rats. PPAR responsiveness of these
genes in vivo in mice has also been modeled with etomoxir, which
inhibits CPT I and is presumed to activate PPAR
indirectly by
causing accumulation of cellular FA (2, 10). Further
evidence for the importance of PPAR
regulation of some of these
genes was provided by the absence of their regulation in PPAR
knockout mice (2). These studies, in combination with the
CPT I upregulation by UF-5 in vivo, suggest that stimulation of
mitochondrial FA
-oxidation represents an additional TG-lowering
mechanism of PPAR
agonists. Consistent with this, UF-5 markedly
stimulated palmitate
-oxidation in human hepatic and skeletal muscle
cells in the present study. Although UF-5 is a potent activator of
human PPAR
(4), the effects of UF-5 are not likely due
to PPAR
activation, because PPAR
immunoreactive protein was
undetectable in these cells. The reason for the high (micromolar-range)
concentrations necessary to achieve this relative to PPAR
activation
potency (~50 nM for human PPAR
) is not clear. However, a similar
dissociation between apparent activation potency and functional effects
in untransfected cells has been observed for fenofibrate effects on
gene regulation (28, 30).
In this study, hamster soleus muscle CPT I mRNA levels were increased
2.3-fold in hamsters treated for 3 wk with rising doses of up to 30 mg/kg UF-5, but they were unaffected in animals treated with 30 mg/kg
UF-5 for only 3 days. This result suggests that, although reduction of
apoC-III and increase of CPT I expression in liver may contribute to
short-term TG lowering in hamsters, stimulation of muscle FA
-oxidation does not. However, the data suggest that stimulation of
muscle FA
-oxidation contributes to the greater TG-lowering effect
of 30 mg/kg UF-5 after longer-term administration. Furthermore,
fenofibric acid administration at the doses tested lowered hepatic
apoC-III to a similar extent as did efficacious doses of UF-5 but did
not affect muscle CPT I gene expression and had no TG-lowering
efficacy. These results, by dissociating TG lowering from suppression
of hepatic apoC-III expression, provide further support for the
importance of muscle CPT I upregulation in TG lowering in fat-fed
hamsters. Increased FA
-oxidation did not decrease glucose
utilization in muscle as suggested by the lack of effect of UF-5 on
plasma glucose concentrations. In fact, the decrease in body weight
seen in UF-5-treated hamsters suggests that PPAR
activation
increased energy expenditure in this animal model.
The relative contribution of FA -oxidation and apoC-III regulation
in TG lowering by PPAR
agonists may have implications for
lipoprotein and lipid metabolism. Reduction of apoC-III synthesis, by
stimulating lipoprotein lipase activity, might be expected to increase
the VLDL-TG fractional catabolic rate (1), thereby increasing LDL production and raising LDL levels. In contrast, stimulation of FA
-oxidation would be expected to reduce VLDL-TG production and therefore to lower LDL levels. In fact, UF-5 treatment significantly lowered LDL cholesterol by over 50%, consistent with the
importance of the latter mechanism for TG lowering.
An additional effect of UF-5 in hamsters was a large increase in liver
weight (Tables 1 and 2). This increase probably does not reflect fatty
liver as judged by visual observation. Rather, hepatic hypertrophy
undoubtedly results from peroxisome proliferation (as measured by
increased cyanide-insensitive palmitoyl-CoA oxidation and by an
increased immunoreactive bifunctional enzyme, not shown). In any case,
the observed hepatic hypertrophy raises the possibility that the
TG-lowering effect of UF-5 occurs at least partially through increased
FA utilization in liver for cell membrane lipid synthesis. It is also
possible that the increased hepatic CPT I expression reflects the need
for an increase in cellular ATP production associated with liver
hypertrophy. However, the following observations argue against these
mechanisms: 1) UF-5 stimulated FA -oxidation directly in
liver cells with no associated cellular proliferation or hypertrophy;
2) the TG-lowering effects of less potent, clinically
administered PPAR
agonists such as fenofibrate are dissociated from
liver hypertrophy in humans; and 3) this possibility does
not apply to UF-5-induced increase in muscle FA
-oxidation, because
UF-5 did not cause muscle hypertrophy (as judged by soleus muscle weights).
In summary, the present study supports the hypothesis that a potent
PPAR agonist exerts marked TG-lowering effects. Stimulation of
mitochondrial FA
-oxidation in liver and muscle appears to contribute to maximal hypolipidemic effects of PPAR
activation. These results should be considered in the design and monitoring of
similar pharmacological agents.
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ACKNOWLEDGEMENTS |
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We thank Drs. Linda Merkel and Mark Perrone for helpful discussions; Colleen Charsky, Charles Hanning, and Tambra Peters for excellent technical assistance; and Elizabeth O'Connor, Kevin Darlington, and Henry Sarau for assistance with animal studies. We thank Dr. Ken Page for analysis of UF-5 levels and Dr. Zaid Jayyossi and Margaret Muc for measurements of liver peroxisomal enzymes. We are grateful to Drs. Robert Groneberg for chemical syntheses and Hong Zhu for providing the rat GAPDH probe. We acknowledge MSD Panlabs (Bothell, WA) for plasma HDL and LDL cholesterol determinations.
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FOOTNOTES |
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Present addresses: Anne Minnich and Nian Tian, Division of Respiratory Disease and Rheumatoid Arthritis, Aventis Pharmaceuticals, Bridgewater, NJ 08807; Lisa Byan, Guilford Pharmaceuticals, Baltimore, MD 21205; Glenda Bilder, Valleybrooke Corporate Center, Malvern, PA 19355.
Address for reprint requests and other correspondence: A. Minnich, Aventis Pharmaceuticals, Rt. 202/206, Bridgewater, NJ 08807 USA (E-mail: anne.minnich{at}aventis.com).
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.
Received 19 May 2000; accepted in final form 20 October 2000.
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