Expression of peroxisomal proliferator-activated receptors
and retinoid X receptors in the kidney
Tianxin
Yang1,
Daniel E.
Michele2,
John
Park1,
Ann M.
Smart1,
Zhiwu
Lin1,
Frank C.
Brosius III1,
Jurgen B.
Schnermann2, and
Josephine P.
Briggs1,2,3
Departments of 2 Physiology and
1 Internal Medicine, University of
Michigan, Ann Arbor, Michigan 48109; and
3 National Institute of Diabetes
and Digestive and Kidney Diseases, Bethesda, Maryland
20892
 |
ABSTRACT |
The discovery that
15-deoxy-
12,14-prostaglandin
J2
(15d-PGJ2) is a ligand for the
-isoform of peroxisome proliferator-activated receptor (PPAR)
suggests nuclear signaling by prostaglandins. Studies were undertaken
to determine the nephron localization of PPAR isoforms and their
heterodimer partners, retinoid X receptors (RXR), and to evaluate the
function of this system in the kidney. PPAR
mRNA, determined by
RT-PCR, was found predominately in cortex and further localized to
proximal convoluted tubule (PCT); PPAR
was abundant in renal inner
medulla, localized to inner medullary collecting duct (IMCD) and renal
medullary interstitial cells (RMIC); PPAR
, the ubiquitous form of
PPAR, was abundant in all nephron segments examined. RXR
was
localized to PCT and IMCD, whereas RXR
was expressed in almost all
nephron segments examined. mRNA expression of acyl-CoA synthase (ACS),
a known PPAR target gene, was stimulated in renal cortex of rats fed
with fenofibrate, but the expression was not significantly altered in
either cortex or inner medulla of rats fed with troglitazone. In
cultured RMIC cells, both troglitazone and
15d-PGJ2 significantly inhibited cell proliferation and dramatically altered cell shape by induction of
cell process formation. We conclude that PPAR and RXR isoforms are
expressed in a nephron segment-specific manner, suggesting distinct
functions, with PPAR
being involved in energy metabolism through
regulating ACS in PCT and with PPAR
being involved in modulating
RMIC growth and differentiation.
15-deoxy-
12,14-prostaglandin
J2; reverse
transcription-polymerase chain reaction; acyl-coenzyme A synthase; microdissected nephron segments
 |
INTRODUCTION |
PROSTAGLANDINS (PGs) are known to be important
signaling molecules, formed in response to various extracellular
stimuli by oxygenation and peroxidation of arachidonic acid. After
secretion from cells, PGs act close to the site of formation. A large
body of evidence has established that one mode of action of PGs is initiated by activation of cell surface receptors of the G protein receptor superfamily and subsequent changes in the level of the intracellular messenger molecules cAMP and/or
Ca2+. This classic pathway
probably mediates many of the known biological effects of PGs.
Nevertheless, the complexity of PG signaling has been apparent from the
unique properties of a group of cyclopentenone PGs that contain an
,
-unsaturated carbonyl group in the cyclopentenone ring. Members of this group have been shown to possess
potent cytotoxic activities, the cellular mechanism of which does not seem to be explained by the classic pathway (9). It is known that the A
and J series of PGs can be transported into the nucleus and can
associate with nuclear proteins (24, 25). Recent studies by two
separate groups demonstrate that the terminal metabolite of the J
series of PGs,
15-deoxy-
12,14-PGJ2
(15d-PGJ2), is a ligand for a
nuclear receptor, the
-isoform of peroxisome proliferator-activated
receptor, PPAR
(8, 16). These findings suggest a role for PGs in
PPAR
-mediated nuclear signaling as an alternative to the classic
signaling through PG surface receptors.
PPARs are a group of zinc-finger-containing transcription factors, a
subfamily of the nuclear hormone receptor gene family. To date, three
subtypes of PPARs have been described from several species: PPAR
,
PPAR
(also called PPAR
or NUC-1), and PPAR
(15, 20, 27). Two
PPAR
isoforms,
1 and
2, differing only in a 30 NH2-terminal amino acid
segment, have recently been characterized (23, 34). PPARs
heterodimerize with retinoid X receptor (RXR) and regulate gene
transcription after binding to peroxisome proliferator-responsive elements (PPREs) in the promoter region of target genes (21). Most of
the known target genes of PPARs are involved in lipid metabolism (27).
The kidney is an active site for both production and action of PGs (6,
7), and
12-PGJ2 has been found
in human urine in significant quantities (13). Furthermore, all three
members of the PPAR family are expressed in the kidney (3, 12). It is
possible therefore that J series PGs may act through PPARs and regulate
gene expression in kidney. Thus the present study was undertaken to
determine the nephron localization of PPAR isoforms and their
heterodimer partners, RXRs, and to test for biological activities of
this system in the kidney.
 |
METHODS |
Animals and dissection methods.
Microdissection of nephron segments was performed in kidneys of male
Sprague-Dawley rats (~8 wk of age) as previously described (38). The
following specimens were dissected: glomeruli, proximal convoluted
tubules (PCT), proximal straight tubules, cortical thick ascending
limb, medullary thick ascending limb (MTAL), cortical collecting ducts,
outer medullary collecting ducts, and inner medullary collecting ducts (IMCD). In addition, the macula densa-containing segment (MDCS) was
dissected by separating the terminal portion of the thick ascending
limb, including the macula densa, removing the adherent glomerulus, and
freeing the macula densa as much as possible of adjacent thick
ascending limb cells. In general, 10 glomeruli, 10 MDCS, or 6-10
mm of other tubule segments were dissected and pooled to constitute one sample.
RNA isolation. RNA from glomerular and
tubular samples was isolated as previously described (35). Briefly,
glomerular and tubular samples were thawed in an ice slurry and
sonicated for 15 s. Twenty micrograms of ribosomal RNA from
Escherichia coli (Boehringer Mannheim,
Indianapolis, IN) was added as carrier, and the sample in 100 µl of
guanadine isothiocyanate buffer was layered onto a gradient of cesium
chloride (100 µl of 97% and 20 µl of 40% cesium chloride in 25 mM
sodium acetate buffer) in a 250-µl polycarbonate ultracentrifuge
tube. Samples were centrifuged for 2 h at 300,000 g in an ultracentrifuge (model TLA
100; Beckman Instruments, Fullerton, CA) with a fixed-angle rotor. The
RNA pellet was redissolved in 0.3 M sodium acetate and ethanol precipitated.
Total RNA from rat adult kidney slices and from mouse kidney at various
stages of development was isolated using TRI-Reagent (Molecular
Research Center). Tissue samples were homogenized in TRI-Reagent
solution. After addition of chloroform and centrifugation, the
homogenates separate into three phases: aqueous, interphase, and
organic. RNA was precipitated from the aqueous phase by addition of
isopropanol. Contaminating genomic DNA was removed with RNase-free DNase I (GeneHunter, Brookline, MA). The purified RNA was redissolved in diethyl pyrocarbonate-treated water containing 20 U of RNasin.
RT-PCR. Primer information with
related references is shown in Table 1.
RT-PCR was performed as previously described (35). Briefly, reverse transcription was performed in the presence of 100 U
monkey murine leukemia virus reverse transcriptase (RT) (Superscript; BRL, Gaithersburg, MD), and 0.5 µg
oligo(dT)12-18 (Pharmacia,
Piscataway, NJ). The nucleotide sequences of primers used in these
studies are shown in Table 1. PCR reactions were performed in a total
volume of 50 µl in the presence of 5 pmol of each oligonucleotide
primer, 200 µM dNTP, 10 mM dithiothreitol, 50 mM KCl, 1.5 mM
MgCl2, 10 mM
Tris · HCl, 0.001% gelatin, 1.25 U of
AmpliTaq DNA polymerase, pH 8.3 (Perkin-Elmer Cetus, Norwalk, CT), and 1.5 µCi
[32P]dCTP (Amersham,
Arlington Heights, IL). The samples were first denatured at 94°C
for 3.5 min followed by 32 PCR cycles as follows: 94°C for 1 min
(melt), 58°C for 1 min (anneal), and 72°C for 1 min (extend).
The last cycle was followed by additional extension incubation of 8 min
at 72°C.
Analysis of PCR products. After
amplification, PCR products were subjected to size separation by
polyacrylamide gel electrophoresis. The band intensity was determined
by phosphoimaging with the Phosphor Analyst software on a GS-250
Molecular Imager System (Bio-Rad, Hercules, CA). To confirm the
identity of the PCR products of the PPAR isoforms, the products were
gel purified and directly sequenced using an ABI 373 DNA sequencer.
Cell culture. Characteristics and in
vitro maintenance of renal medullary interstitial cells (RMICs) have
been described previously (18). The present study has demonstrated the
presence of lipid droplets in these cells by oil red O staining method
(data not shown). Troglitazone and
15d-PGJ2 were dissolved in DMSO
with a final concentration of 0.1% DMSO in culture medium. Control medium contained 0.1% DMSO. Cells were plated at a density of 4.3 × 104 cells/ml and were
grown in RPMI medium supplemented with 5% FBS in the presence of 10 µM troglitazone, 1 µM
15d-PGJ2, or 0.1% DMSO. After
incubation, the cells were trypsinized, and the cell number was counted
at the times indicated. To examine the cell morphology, the cells were
directly stained with fluorescein diacetate.
 |
RESULTS |
mRNA expression of PPAR
isoforms in
fat tissues and kidney.
PPAR
is known to have two isoforms, PPAR
1 and PPAR
2, which are
alternatively spliced products of the same gene. PPAR
2 differs from
PPAR
1 by a sequence of 30 additional
NH2-terminal amino acids. Using a
primer pair specific to PPAR
2 (Fig.
1A), the 100-bp PCR product specific for PPAR
2 was detected in fat tissue, but not in kidney, whereas the nonspecific 793-bp product was
detected in both tissues (Fig. 1B),
indicating that the isoform expressed in kidney is PPAR
1 and that
PPAR
2 is absent.

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Fig. 1.
A: primer design for amplification of
two peroxisome proliferator-activated receptor (PPAR) -isoforms.
B: RT-PCR from fat and kidney with two
sets of primers: sense primer 1 and antisense primer 1 (S1 and AS1) are
specific for PPAR 2, and sense primer 2 and antisense primer 2 (S2
and AS2) are common to both PPAR 1 and - 2
isoforms.
|
|
Distribution of PPAR subtype mRNA in kidney
regions. To amplify PPAR
, -
, and -
, primers
were selected in divergent regions of the published cDNA sequences
(Table 1). As shown in Fig. 2, products of
expected size were obtained for the PPAR isoforms (523 bp for PPAR
,
496 bp for PPAR
, and 793 bp for PPAR
). PPAR
mRNA was present
in cortex, PPAR
mRNA was present in inner medulla, and PPAR
mRNA
was present in both cortex and inner medulla. Product identity was
confirmed by direct sequencing. When PCR for PPAR isoforms was
performed in the absence of reverse transcription, there was no
recognizable band, indicating the origination of the products from
mRNA, not from genomic DNA (data not shown).

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Fig. 2.
Distribution of mRNAs of three PPAR subtypes ( , , and ) in the
kidney regions by RT-PCR. PCR products were separated on 5%
polyacrylamide gel and analyzed by phosphoimage.
|
|
Nephron localization of mRNA of PPAR subtypes.
RT-PCR for each PPAR isoform was performed on cDNA derived from
microdissected nephron segments from 10-wk-old Sprague-Dawley rats. All
three isoforms were tested in each sample, with four sets of
determinations performed to localize isoform expression. Figure
3 shows a representative gel. Each isoform
exhibited a unique and consistent distribution pattern. PPAR
mRNA
was detected in PCT, PPAR
mRNA was found in IMCD, and PPAR
mRNA
was found in all nephron segments examined. RT-PCR of
-actin was run
to verify homogeneity of mRNA amounts in each sample. A single 351-bp
band for
-actin with similar intensity was detected in all samples.

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Fig. 3.
Localization of mRNA expression of three PPAR subtypes ( , , and
) in isolated nephron segments of Sprague-Dawley rats. PCR for the
three PPAR subtypes and -actin was performed on the same cDNA
(representative example from 4 separate experiments). IMCD and OMCD,
inner and outer medullary collecting ducts, respectively; CCD, cortical
collecting duct; MDCS, macula densa-containing segment; CTAL and MTAL,
cortical and medullary thick ascending limbs, respectively; PST and
PCT, proximal straight and convoluted tubules, respectively; and Glm,
glomeruli.
|
|
Nephron localization of mRNA of RXR subtypes.
Since RXR is the required heterodimer partner of PPAR, we also examined
the nephron localization of two isoforms of RXR, RXR
and RXR
. As shown in Fig. 4, RXR
mRNA was present
predominately in PCT and IMCD, whereas RXR
was present in all
nephron segments examined except MTAL.

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Fig. 4.
Localization of mRNA expression of two retinoid X receptor (RXR)
subtypes ( and ) in isolated nephron segments of Sprague-Dawley
rats.
|
|
mRNA expression of PPAR
in cultured
kidney cell lines.
RT-PCR for PPAR
was performed on cDNA derived from cultured kidney
cell lines including M-1 (30), mIMCD-K2 (32), and RMIC (18). PPAR
mRNA was detected in all three cell lines tested, with relatively
abundant expression in RMIC (Fig. 5).

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Fig. 5.
mRNA expression of PPAR in cultured cells of kidney origin.
Representative gel shows PCR product of PPAR from RMIC, mIMDC-K2,
and M1 cells which were derived from renal medullary interstitial
cells, mouse inner medullary collecting duct cells, and mouse cortical
collecting duct cells, respectively.
|
|
mRNA expression of PPAR during mouse kidney
development. RT-PCR for the three PPAR subtypes was
performed on cDNA derived from the developing mouse kidney beginning at
embryonic day E14.5 to 2 wk after
birth. As shown in Fig. 6, PPAR
mRNA
expression was increased in late gestation period and tended to
decrease after birth. In contrast, PPAR
and PPAR
mRNA expression
gradually increased with development, with high levels maintained after birth.

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Fig. 6.
mRNA expression of three PPAR subtypes ( , , and ) during mouse
kidney development. Whole kidney was dissected from different
developmental periods, and PCR was performed on 10× and
100× dilutions of cDNA. E, embryonic; PND, postnatal
developmental.
|
|
Activation of gene expression by PPAR
ligands. Acyl-CoA synthase (ACS) is a well-established
target gene of PPARs. To evaluate the functional role of PPARs in the
kidney, we examined the effect of fenofibrate and troglitazone,
activators for PPAR
and PPAR
(2), respectively, on renal ACS mRNA
expression. ACS mRNA in renal cortex was significantly stimulated in
animals fed with fenofibrate (0.5% in food) (28) for 1 wk, but
medullary expression was unchanged, consistent with the distribution
pattern of PPAR
in the kidney (Fig. 7,
left). As expected, treatment with
fenofibrate also significantly induced ACS mRNA expression in liver
(Fig. 7, right), another major site
of PPAR
expression. In contrast, administration of troglitazone by
gavage (100-400
mg · kg
1 · day
1)
for 3 days had no obvious effect on ACS mRNA expression in either cortex or medulla (data not shown).

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Fig. 7.
Regulation of mRNA expression of acyl-CoA synthase (ACS) in kidney and
liver in vivo by fenofibrate (F). Sprague-Dawley rats were fed with
fenofibrate for 1 wk, and RT-PCR for ACS and -actin were performed
on 1 µg RNA isolated from kidney regions and liver.
|
|
Effect of PPAR
on RMIC growth and
differentiation.
PPAR
has been implicated in modulation of cell growth and
differentiation in PPAR
-expressing cells. Therefore, we examined the
effect of PPAR
ligands on the cell growth in RMIC. Proliferating RMICs were treated with 10 µM troglitazone, 1 µM
15d-PGJ2, or 0.1% DMSO for the
times indicated. As shown in Fig. 8, both
troglitazone and 15d-PGJ2
significantly inhibited cell growth, and
15d-PGJ2 was more potent than
troglitazone in growth inhibitory activity (Fig. 8). PPAR
ligands
also induced dramatic changes in cell morphology in RMIC characterized
by extensive process formation (Fig. 9).

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Fig. 8.
Effect of PPAR ligands on cell growth of RMIC. Cells were plated at
a density of 4.3 × 104
cells/ml and exposed to 10 µM troglitazone, 1 µM
15-deoxy- 12,14-prostaglandin
J2
(15d-PGJ2), or 0.1% DMSO. Cells
were trypsinized and counted at the times indicated.
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Fig. 9.
Effect of PPAR ligands on morphology of cultured RMIC. RMICs were
grown and treated as described in Fig. 8. Cells were stained directly
with fluorescein diacetate after treatment for 3 days.
|
|
 |
DISCUSSION |
Over the last decade and a half, a large number of putative hormonal
nuclear receptors have been identified, but for many, including those
in the PPAR family, the natural ligands have been unknown, and
therefore their physiological role has been uncertain (21). Recently,
there has been remarkable progress in the identification of natural
ligands of this nuclear receptor subclass, with
15d-PGJ2 identified as a ligand of
PPAR
(8, 16) and leukotriene B4 identified as a natural ligand for PPAR
(4). In the present studies,
we examined the renal distribution of expression of PPAR
, -
, and
-
, and their obligate heterodimer RXR partners and show that ligands
for the PPAR receptors alter gene expression or cell viability in a
pattern consistent with the distribution of their receptors. These
studies support the hypothesis that PGs, including 15d-PGJ2, may act through the PPAR
receptor class to regulate gene expression and cell growth in the kidney.
The PPAR subfamily consists of three members, PPAR
, -
, and -
,
which share a high degree of similarity in their overall amino acid
sequences, particularly in the DNA binding domain (5). The three PPARs
bind to the same PPRE in the promoter regions of their target genes and
appear to have similar effects on gene transcription of several enzymes
involved in fatty acid oxidation in vitro (27). They bind many of the
same ligands, although with variable affinities (27). Distinct
functions for PPAR family members are suggested from their
tissue-specific expression patterns: PPAR
mRNA is mainly expressed
in liver, kidney, and heart; PPAR
is mainly expressed in fat tissues
and spleen; PPAR
, which is also called PPAR
or NUC-1, is
expressed widely (3, 17).
PPAR activators are known to produce peroxisome proliferation in
kidney, but the kidney is a highly heterogeneous organ, with multiple
distinct cell types and functions. Study of gene expression of
different subtypes of PPARs in defined nephron segments is one approach
by which to study their cell type-specific functions in the kidney. Our
data demonstrated that three PPARs had distinct localization patterns
in kidney regions and in isolated nephron segments. PPAR
mRNA was
expressed predominately in cortex and further localized to PCT,
consistent with previous reports (3, 12). PPAR
was expressed only in
renal medulla, but not in cortex, and further localized to IMCD and
RMIC. This finding is largely but not completely in agreement with the
study by Guan et al. (12) in which PPAR
message was
found to be present in both cortex and medulla and was localized
predominantly in IMCD but not in RMIC. The discrepancy may be due to
differences in species or in experimental protocols. We found that
PPAR
was detected in both cortex and medulla and was expressed in
all nephron segments examined, consistent with the observation by in
situ hybridization technique (3) and with the ubiquitous expression
noted previously (17).
PPAR requires RXR as a heterodimer partner to regulate PPAR target gene
transcription. In vitro studies have shown that all three PPAR subtypes
can interact with either RXR
, -
, or -
(17, 19). However, we
found that RXR
mRNA was specifically colocalized with PPAR
and
PPAR
in PCT and IMCD, respectively, favoring the occurrence of
heterodimers of PPAR
-RXR
in PCT and PPAR
-RXR
in IMCD.
RXR
was expressed in all nephron segments, coinciding with the
expression pattern of PPAR
, suggesting that both receptors may have
a constitutive function for all nephron segments. The partnership
between PPAR and RXR in vivo will be complex since RXR can also serve
as the heterodimer partner for other hormones such as thyroid hormone
and vitamin D receptors.
PPAR
, the first member of the PPAR subfamily, was cloned as an
orphan nuclear receptor activated by agents that induce peroxisome proliferation (14). There is accumulating evidence that PPAR
is a
major mediator for regulation of energy homeostasis. PPAR
mRNA is
predominantly distributed in tissues capable of oxidizing fatty acids, such as liver, kidney, and heart. In extension of the
initial observation by Braissant et al. (3), we showed that PPAR
mRNA was localized in renal cortex and in proximal tubules
where high peroxisomal
-oxidation activity has been described (26).
In support of these findings, we showed that ACS, a known PPAR target
gene, was stimulated in renal cortex but not medulla of rats fed with
fenofibrate, an efficient activator for PPAR
. Indeed, transport by
the proximal tubule is among the highest of all nephron segments, and
as a consequence this segment is highly vulnerable to damage caused by
energy depletion. Our data suggest that PPAR
may be involved in
energy supply to proximal tubule through regulation of gene
transcription of such PPAR target genes as ACS. Although in vitro
studies suggest that all PPAR isoforms can regulate transcription of
the group of genes involved in fatty acid oxidation, we found that in
vivo renal ACS is regulated by fenofibrate but not by troglitazone.
This represents another line of evidence for distinct function of PPAR
isoforms in vivo. PPAR
is probably involved in energy metabolism in
PCT as discussed above, whereas the abundant expression of PPAR
in
renal medulla and RMICs suggest that in these cells it may serve other functions.
The J series of PGs are known to possess potent antitumor,
antiproliferation, and antivirus activity, but the biological
significance and the cellular mechanism underlying these effects are
poorly understood (9). The discovery that
15d-PGJ2 is a ligand for PPAR
suggests that this receptor may mediate the cytotoxic effect of the J
series of PGs. This hypothesis has been supported by recent evidence.
Thiazolidinedones have been shown to inhibit cell growth
in cultured human aorta and coronary myocytes (22). More recently,
ectopically expressed PPAR
has been shown to induce cell cycle
withdrawal in cultured NIH-3T3 cells, in addition to its effect in
inducing adipogenesis (1). The present study shows that both
troglitazone and 15d-PGJ2 have a
similar effect in inducing growth inhibition and differentiation of
RMICs. These findings suggest a possibility that PPAR
may be the
major receptor mediating the wide range of biological activities of the
J series of PGs. This issue needs to be more extensively investigated
in future studies.
Previous studies, using whole embryo and adult tissues, have shown that
the three PPAR subtypes are differentially expressed during development
(17). To address this issue in the kidney, we examined expression
patterns of the three subtypes during murine kidney development. All
three PPARs were expressed in early embryonic kidneys; PPAR
and
PPAR
mRNA levels increased with development and remained elevated 2 wk after birth, whereas PPAR
mRNA peaked in late gestation. These
data demonstrate that the three subtypes of PPARs are differentially
regulated during murine kidney development, providing additional
evidence to support the distinct function of PPAR
subtypes in renal function.
In summary, different nephron localization patterns of various subtypes
of PPAR/RXR suggest their distinct functions in different nephron
segments. PPAR
is localized to renal cortex and further localized to
PCT; activation of PPAR
by fenofibrate stimulates ACS gene
expression in cortex, but not renal medulla, consistent with the
distribution pattern of PPAR
and suggesting a role of PPAR
in
energy metabolism in PCT. However, PPAR
is abundant in inner medulla
and localized in IMCD and RMIC. Activation of PPAR
induces RMIC
growth inhibition and cell shape changes, suggesting a potential role
of PPAR
in controlling the proliferation and differentiation of RMIC.
 |
ACKNOWLEDGEMENTS |
We are grateful to Drs. E. Nord, G. Fejes-Toth, and B. Stanton for
generously supplying us with renal medullary interstitial cells, M-1
cells, and mIMCD-K2 cells. The gift of troglitazone from Dr. J. W. Johnson (Parke-Davis) is gratefully acknowledged.
 |
FOOTNOTES |
This work was supported by National Institute of Diabetes and Digestive
and Kidney Diseases Grants DK-37448, DK-39255, and DK-40042. Additional
support was in part from the General Clinical Research Center at the
University of Michigan, funded by National Center for Research
Resources Grant M01-RR-00042.
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: J. P. Briggs,
NIDDK, NIH, Bldg. 31, Rm. 9A17, 31 Center Drive MSC 2560, Bethesda, MD
20892 (E-mail: BriggsJ{at}hq.niddk.nih.gov).
Received 24 August 1998; accepted in final form 1 July 1999.
 |
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