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Women’s Health Research Institute, Wyeth-Ayerst Research, Radnor, Pennsylvania 19087
Address all correspondence and requests for reprints to: Dr. Heather A. Harris, Women’s Health Research Institute, Wyeth-Ayerst Research, 145 King of Prussia Road, Radnor, Pennsylvania 19087. E-mail: harrish{at}war.wyeth.com
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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or ERß. One gene, metallothionein II, was regulated by
both receptor subtypes in LNCaPLN3 cells, but only by ERß in SAOS-2
cells. Because cycloheximide blocks this response, induction is
probably mediated through regulation of at least one other protein.
Identification of endogenous genes that are regulated differentially by
ER and ERß will be valuable tools in elucidating the function of
ERß and the mechanisms by which these two receptors regulate
transcription. |
Introduction |
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. Since this initial report,
ERß has been cloned from a number of other species, including human,
mouse, cow, and goldfish (2, 3, 4, 5, 6, 7). After the discovery of
ERß, most work has focused on mapping the distribution of its
messenger RNA (mRNA) in normal and neoplastic tissues, characterizing
its binding affinity for and trans-activational activity
with a wide variety of ligands, and assessing its interaction with
ER.
ERß mRNA is detectable by RT-PCR and in situ hybridization
in a wide variety of tissues and has been best mapped in the rat and
mouse. Its distribution is largely distinct from that of ER,
although receptors are coexpressed in some tissues. For example, ERß
is the predominant receptor subtype in ovarian granulosa cells
(8, 9), prostate epithelial cells (8),
bladder (10), and paraventricular nucleus and cortex of
the brain (11, 12). The rat uterus expresses primarily
ER (13, 14), whereas both receptors are detected in
bone (15, 16, 17) and the cardiovascular system (18, 19). Because of its pattern of distribution, especially its lack
of expression in the rodent uterus, ERß has potential as an
attractive drug target (20). Development of
ERß-selective ligands will test the hypothesis that
receptor-selective hormone replacement therapy will retain many
benefits of traditional replacement therapy but reduce the concern over
estrogen’s effects in the uterus. In addition, because of its high
expression in the rodent prostate, an ERß-selective ligand may have
therapeutic value in males.
Although the ligand-binding domains of human ER and ERß are only
59% identical at the amino acid level (21), the
affinities of binding 17ß-estradiol are quite similar. However, other
compounds show marked selectivity for either ER or ERß. Genistein
is a phytoestrogen that binds with approximately 10- to 37-fold higher
affinity for ERß (1, 22, 23). On the other hand,
16-iodo-17ß-estradiol, a common iodinated radioligand for ER
studies, is ER selective (24). Thus, partially
selective ligands already exist, and the question remains of whether
this selectivity can be improved to the point where only one receptor
subtype is activated.
Despite the abundance of data characterizing ERß and much speculation
on the subject, its function is essentially unknown. Knockout mice are
available to help elucidate the functions of ER and ERß. ER
knockout (ERKO) mice lack ER (25) and have a plethora
of defects, primarily centering on reproductive system function
(reviewed in Ref. 26). Intriguingly, however, some
estrogenic responses remain in these mice. For example, one study
compared the effectiveness of estrogen treatment in ameliorating
consequences of artificially induced vascular injury to carotid
arteries in wild-type and ERKO mice (27). In both types of
mice, pharmacological doses of estradiol suppressed the increase in
medial area and smooth muscle proliferation seen in the vehicle-treated
animals. It is thought that these responses to endothelial denudation
may narrow the lumen of the vessel, thus restricting blood flow.
Because estradiol was equally effective in ERKO as wild-type mice, one
interpretation is that ER is not necessary for this response.
Because ERß mRNA is also expressed in these vessels, perhaps it
mediates the action of estrogen. However, direct evidence supporting
this hypothesis is lacking. In fact, in the ERßKO mice, estradiol
remains protective, and data from the double ERKO mice are needed to
settle the question of whether this activity can be mediated via ER
or ERß (28).
Two other reports suggest a function for ERß in the cardiovascular system. First, ERß mRNA is strongly up-regulated in injured rat aortas and carotid arteries (19, 29). Second, using a carotid denudation model, Makela et al. (29) show that genistein is as potent and efficacious as 17ß-estradiol in reducing intimal cell proliferation in ovariectomized rats, but, unlike 17ß-estradiol, it does not increase uterine wet weight.
Mice lacking ERß have only recently been described and are not fully characterized (30). However, ovulation is reduced, and it has been suggested that bladder and prostate hyperplasia occurs as the animals age.
We have taken a different approach toward identifying the function of
ERß. Using a differential display strategy, mRNA populations of two
cell lines were compared before and after treatment with
17ß-estradiol. We chose cell lines normally expressing ERß, but not
ER, mRNA as determined by RT-PCR. Before treatment with estradiol,
cells were engineered to overexpress ERß to boost the magnitude of
the response. This report focuses on one gene that is selectively
regulated by the two ERs: metallothionein II.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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E1a vector
plasmid. This plasmid contained adenovirus sequences from map unit
0–17, with a deletion of E1a region between map units 1.4–9.1. The
hERßL transcription unit in Ad5E1a plasmid
contained the cytomegalovirus 1E promoter, Ad5 tripartite leader,
coding sequence hERßL, and simian virus 40 late
polyadenylation signal sequences. hERßL in Ad5
E1a plasmid was then linearized and transfected along with the
ClaIA fragment of Ad5 virus with the E3 region deletion
(80–88 map units) into 293 cells. Viral plaques generated by
homologous recombination were isolated, amplified, and characterized by
Hirt DNA analysis and cell lysis assay in A549 cells. All of the above
tests indicated that the recombinant Ad5 hERßL
virus contained the expected DNA fragments and was replication
defective. The virus was further purified by replaquing. The isolated
plaques were amplified, tested, and used as a seed stock to generate
large amounts of the virus in 293 cells. The virus was titrated in 293
cells by plaque assay, and the stock contained 1.28 x
109 plaque-forming units/ml.
Evaluation of SAOS-2 and LNCaPLN3 cells for endogenous ER mRNA
Total RNA was isolated from LNCaPLN3 and SAOS-2 cells using
TRIzol (Life Technologies, Inc., Gaithersburg, MD)
according to the manufacturer’s directions. The samples were treated
with ribonuclease-free deoxyribonuclease I (Life Technologies, Inc.) at 1 U/µg for 30 min at 37 C. RNA was purified using
RNeasy columns (QIAGEN, Valencia, CA), and the
concentration was estimated by UV spectrophotometry.
RT reactions were performed on 0.5 µg RNA in a 20-µl reaction. For
ER the reaction contained 1x PCR buffer (Life Technologies, Inc.), 5 mM MgCl2, 1.25
µM ER-specific reverse primer
(5'-CCAGCAGCATGTCGAAGATC-3'), 0.5 µM
glyceraldehyde-3-phosphate dehydrogenase (GAPDH)-specific reverse
primer (5'-CACCCTGTTGCTGTAGCCAAATTC-3'), 0.5 mM deoxy
(d)-NTPs, 20 U RNasin (Promega Corp., Madison, WI), and
200 U Superscript II reverse transcriptase (Life Technologies, Inc.). The ERß reaction contained the same components as ER
with the exception of 2.5 mM MgCl2
and 1.25 µM ERß-specific reverse primer
(5'-GCAGAAGTGAGCATCCCTCTTTG-3'). A duplicate reaction that was
identical in all reagents except that it did not contain Superscript II
reverse transcriptase was performed for each sample as a negative
control to ensure that the RNA samples were not contaminated by DNA.
Reactions were incubated at 42 C for 15 min, followed by 5 min at 99 C
and 5 min on ice before amplification.
PCR was initiated by adding 80 µl of a master mix containing
ER-specific forward primer (5'-GGAGACATGAGAGCTGCCAAC-3') or
ERß-specific forward primer (5'-CAGCATTCCCAGCAATGTCAC-3') and
GAPDH-specific forward primer (5'-GACATCAAGAAGGTGGTGAAGCAG-3') directly
to the 20-µl RT reaction. The final concentrations of reagents in the
ER 100-µl PCR reaction were as follows: 0.25 µM of
each ER-specific primer, 0.1 µM of each GAPDH primer,
1 x PCR buffer (Life Technologies, Inc.), 0.2
mM dNTPs, 2 mM MgCl2, and
0.5 U Taq DNA polymerase (Life Technologies, Inc.). The ERß reaction contained the same amount of reagents,
except for 1 mM MgCl2.
Two-step PCR was carried out in a PE 9600 for 25 cycles as follows: 95
C for 30 sec and 64 C for 1.5 min. Samples were incubated at 64 C for
10 min after amplification.
Twenty microliters of each sample were separated using a 1.5% agarose
gel and transferred to Hybond-N+ (Amersham Pharmacia Biotech, Piscataway, NJ) by alkali Southern blotting
in 0.4 N NaOH and 0.6 M NaCl. Blots were
prehybridized at 42 C for 30 min in Rapid-Hyb buffer (Amersham Pharmacia Biotech). Oligonucleotide probes specific for ER
(5'-TGAACCAGCTCCCTGTCTGCCAGGTTGGT-3'), ERß
(5'-CCGCATACAGATGTGATAACTGGCGATGGA-3'), and GAPDH
(5'-GCTGTTGAAGTCACAGGAGACAACCTGGT-3') fragments were end labeled with
[
-32P]ATP using polynucleotide inase
(Life Technologies, Inc.). Probes were added to the blot
at 3.0 x 106 cpm/ml and incubated at 42 C
for 1 h. ER and GAPDH hybridizations were performed independently.
Blots were washed once in 2 x SSC (standard saline citrate) and
0.1% SDS at room temperature for 15 min, then twice in 0.2 x SSC
and 0.1% SDS at 42 C for 15 min. Blots were then exposed to film.
Cell culture, infection, and treatment
All tissue culture reagents were purchased from Life Technologies, Inc. SAOS-2 cells were maintained in monolayer
culture using McCoy’s 5A medium supplemented with 10% FBS, 2
mM GlutaMAX-1, 100 U/ml penicillin G, and 100 µg/ml
streptomycin sulfate. Sixteen hours before infection, the cells were
plated in phenol red-free RPMI 1640 medium supplemented with 10%
charcoal/dextran-treated (stripped) FBS (HyClone Laboratories, Inc., Logan, UT), 2 mM GlutaMAX-1, 100 U/ml
penicillin G, and 100 µg/ml streptomycin sulfate. This medium was
used for the remainder of the experiment. Cells were infected with a
1:20 dilution of Ad5 hERßL virus or Ad5 hER
virus, or using 2% stripped FBS in phenol red-free medium with
antibiotics and GlutaMAX-1 for 2 h at 37 C. Medium containing
virus was aspirated, and the cells were washed with medium. Fresh
medium was added, and the cells were allowed to recover overnight at 37
C. Cells were then treated with various compounds for 24 h. The
17ß-estradiol and diethylstilbestrol (DES) were added at 10
nM, genistein was given at 0.1 µM, and
ICI-182780 and RU486 were given at 1 µM. Test compounds
were each given at 1 µM. Total RNA was isolated using the
TRIzol method (Life Technologies, Inc.). As a positive
control for ER function, SAOS-2 cells were coinfected with Ad5-ER
and Ad5 2 x estrogen response element (ERE)-thymidine kinase
(TK)-luciferase viruses, then treated with various concentrations of
17ß-estradiol with or without 1 µM ICI-182780, and
luciferase activity was measured.
For the cell line survey of metallothionein II expression, all cell lines were plated, infected, and treated as described above. However, lines were grown in the following media supplemented with 10% stripped serum, 2 mM GlutaMAX-1, 100 U/ml penicillin, 100 µg/ml streptomycin sulfate: Hep-G2, DMEM and 1 x nonessential amino acids; Ishikawa, DMEM/Ham’s F-12 (1:1 mix); MCF-7, DMEM/Ham’s F-12 (1:1 mix) and 1 mM sodium pyruvate; human aortic endothelial cells, endothelial basal medium and 0.1% antibiotic/antimycotic; and D-12, DMEM/Ham’s F-12 (1:1 mix) and serum that was not heat inactivated. All cell lines were switched to phenol red-free medium containing 2% stripped serum at infection as described above, except MCF-7, which was infected and treated in 10% stripped serum.
Rapid analysis of differential expression (RADE)
RT. After deoxyribonuclease I treatment, 6 µg total RNA
were incubated with 1 x RT buffer [25 mM Tris-Cl (pH
8.3), 37.6 mM KCl, 3 mM
MgCl2, and 5 mM dithithreitol, from
Genhunter, Nashville TN), 20 µM dNTPs; Life Technologies, Inc.), and 0.2 µM
HT11C (oligonucleotide AAGCTTTTTTTTTTTC)
in a final volume of 600 µl. This reaction mixture was incubated at
65 C for 5 min to denature secondary structures, followed by a 10-min
incubation at 37 C. At this time 30 µl Superscript II reverse
transcriptase (200 U/µl; Life Technologies, Inc.) were
added to the reaction, and incubation proceeded for 1 h at 37 C.
The enzyme was inactivated by heating at 75 C for 5 min. An aliquot of
this reaction was then used for the second strand synthesis by PCR.
PCR
Two microliters of the RT reaction were used for PCR, and the
final concentrations of reagents were 10 mM Tris-Cl (pH
8.4), 100 mM KCl, 1.5 mM
MgCl2 and 0.001% gelatin, 2 µM
dNTPs, 15 nM [33P]dATP (NEN Life Science Products, Boston, MA), 1 U AmpliTaq DNA polymerase
(Perkin-Elmer Corp., Norwalk, CT), and 1 µM
arbitrary primer (5'-AAGCTTGCCATGG-3') for a total reaction volume of
20 µl. This reaction mixture was then thermocycled using the
following parameters: 92 C for 2 min, one cycle; 92 C for 15 sec, 40 C
for 2 min, and 72 C for 30 sec, 40 cycles; and 72 C for 5 min.
Gel electrophoresis
PCR products were separated by gel electrophoresis on a 6%
denaturing polyacrylamide gel (5.7% acrylamide, 0.3% bisacrylamide,
42% urea, and 51% H2O) in 1 x TBE buffer
(0.1 M Tris, 0.09 M boric acid, and 1
mM EDTA) for 3 h at 2000 V. The gel was then
transferred to filter paper (Schleicher & Schuell, Inc.,
Keene, NH), dried under vacuum at 80 C for 1 h, and exposed to
x-ray film for 24 h. The developed film was then superimposed over
the dried gel, and the band of interest was identified. Band corners
were marked using a 22-gauge syringe needle, and the gel slice within
these boundaries was excised with a razor blade and immersed in 100
µl H2O. The sample was boiled in a water bath
for 15 min and centrifuged for 2 min, and the supernatant solution was
transferred to a new tube. Added to this sample were 5 µl 10 mg/ml
glycogen, 10 µl 3 M sodium acetate, and 450 µl 100%
ethanol. The sample was mixed, allowed to precipitate overnight at -20
C, and centrifuged for 10 min at 10,000 x g. The
supernatant solution was removed, and the pellet was washed with 200
µl 85% ethanol, dried, and resuspended in 10 µl
H2O. A 3-µl aliquot of this was used in a
reamplification PCR reaction in the presence of 1 x PCR buffer,
20 µM dNTPs, 0.2 µM
arbitrary primer, 0.2 µM oligonucleotide
HT11C, and 2 U AmpliTaq polymerase using the same
cycling parameters as the PCR reaction above. The resulting product was
then used as probe in a Northern hybridization assay to confirm
regulation and was cloned into a bacterial plasmid for sequence
analysis.
Fragment cloning
Reamplified RADE fragments were cloned into pCR2.1 using the TA
cloning kit (Invitrogen, San Diego, CA). After lysis,
colonies were screened by PCR for the correct insert size. Colonies
were lysed in 20 mM Tris-HCl (pH 8), 50 mM KCl,
2.5 mM MgCl2, 0.5% Tween-20, and 100
µg/ml proteinase K by incubating for 30 min at 56 C, then for 10 min
at 99 C to inactivate the proteinase K. Of this reaction 2 µl were
used in a PCR reaction of 20 mM Tris-HCl (pH 8), 50
mM KCl, 2.5 mM MgCl2, 75
µM dNTPs, 375 nM M-13 forward and reverse
primers, and 2.5 U Taq polymerase (Life Technologies, Inc.). The reactions were cycled as follows: 95 C for 30 sec, 64
C for 30 sec, and 72 C for 45 sec for 30 cycles.
Fragment sequencing
Clone 6a.2 was sequenced according to the ABI Prism Dye
Terminator Cycle Sequencing Ready Reaction Kit with AmpliTaq DNA
polymerase using the recommended protocol from PE Applied Biosystems (Foster City, CA). Spin columns (AGTC) were employed
to remove unincorporated dye-labeled nucleotides after cycle
sequencing. Automated DNA sequencing grade 4.75% polyacrylamide gels
were run for all of the DNA sequencing samples using ABI 373 DNA
sequencers. Sequencing data were edited using Sequence Navigator
(PE Applied Biosystems) and were assembled using either
DNAStar (DNAStar, Inc., Madison, WI) or Sequencher 3.0 (Gene
Codes Corp., Ann Arbor, MI). The sequence was trimmed of RADE
primers and used in a BLAST search using Millennium software.
Isolation of cDNA probe from clone 6a.2
The insert from clone 6a.2 was isolated from the plasmid using
PCR as described above for colony screening. The PCR product was
purified from an agarose gel using Wizard Preps (Promega Corp.).
Northern blot analysis of regulation
Polyadenylated RNA was isolated from total RNA using an Oligotex
mRNA isolation kit (QIAGEN) according to the
manufacturer’s instructions. Six micrograms of mRNA or 10 µg total
RNA were separated on a 1.5% agarose, 0.22 M formaldehyde,
10 mM HEPES, and 1 mM EDTA gel. RNA was
transferred to Hybond-N (Amersham Pharmacia Biotech) by
capillary action in 20 x SSC (standard saline citrate) overnight.
After transfer, the membrane was UV cross-linked and dried at 80 C for
10 min. Northern blots were prehybridized in Rapid Hyb solution
(Amersham Pharmacia Biotech) for 30 min. Reamplified RADE
fragments for band 6a or fragment from clone 6a.2 were random primer
labeled using the Redi-Prime kit (Amersham Pharmacia Biotech) according to the manufacturer’s instructions.
Unincorporated nucleotides were removed using a Nap-5 column
(Amersham Pharmacia Biotech), and incorporation of
[32P]dCTP was measured by liquid scintillation
counting. The probes were denatured at 100 C for 10 min, and 1.5
x 106 cpm/ml Rapid-Hyb hybridization solution
were added to the membrane. The blots were hybridized at 65 C for
5 h and were washed as follows: once in 2 x SSC and 0.1%
SDS at 65 C for 15 min, and twice in 0.2 x SSC and 0.1% SDS at
65 C for 15–30 min. Blots were exposed to film and to a PhosphorImager
screen (Molecular Dynamics, Inc., Sunnyvale, CA). After
probing with 6a RADE fragment or 6a.2 cloned fragment, the blot was
probed with a cDNA homologous to GAPDH as described above.
Hybridization signal from 6a fragment was normalized to that of GAPDH
on a PhosphorImager (Molecular Dynamics, Inc.) to
determine the fold induction.
Treatment of SAOS-2 cells with cycloheximide
Verifying the effect of cycloheximide on protein synthesis.
Cells were plated as described above, infected with hERß long virus
for 2 h, and allowed to recover overnight. Cells were then treated
with 10 µg/ml cycloheximide for 1 h at 37 C. After this
incubation, medium was aspirated and replaced with medium containing 10
µg/ml cycloheximide and 50 µCi/ml
[35S]methionine (NEN Life Science Products) in methionine-deficient medium with or without 10
nM 17ß-estradiol, and incubation was continued at 37 C
for 8 or 24 h. Control cells were not treated with 17ß-estradiol
or cycloheximide. Protein was harvested from cells by washing plates
three times in cold PBS, then scraping cells off plates in 500 µl
PBS. Cells were pelleted and resuspended in 200 µl RIPA buffer (150
mM NaCl, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS,
and 50 mM Tris, pH 8.0). Methionine incorporation was
measured by trichloroacetic acid precipitation. Five and 10 µl of
each sample were spotted on individual Whatman filters
(Clifton, NJ). Filters were boiled in 10% trichloroacetic acid for 10
min, then washed three times for 10 min each time in deionized water,
three times for 10 min each time in 95% ethanol, and once briefly in
acetone. The dried filters were placed in scintillation fluid and
counted for 1 min.
Evaluation of the effect of cycloheximide on 17ß-estradiol
regulation of metallothionein II mRNA
Cells were plated as described above, infected with hERß long
virus for 2 h, and allowed to recover overnight. Control cells
were treated with vehicle or 10 nM 17ß-estradiol for 8 or
24 h. Other cells were treated with 10 µg/ml cycloheximide for
1 h at 37 C. After this incubation, medium was replaced with
medium containing 10 µg/ml cycloheximide with or without 10
nM 17ß-estradiol, and incubation was continued at 37 C
for 8 or 24 h. Total RNA was prepared and analyzed by Northern
blot (described above) or quantitative RT-PCR (qRT-PCR;
described below).
Estimation of the amount of ERß protein in cycloheximide-treated
samples
Duplicate cultures were prepared and treated as described in the
previous section. After the 8-h incubation, cells were washed four
times with DMEM to remove estradiol. Cycloheximide (10 µg/ml) was
added to all samples as well as 1 nM
[3H]estradiol. Some cells were cotreated with
0.3 µM DES to estimate nonspecific binding. After
incubation for 2.5 h at 37 C, cells were washed with DMEM and
lysed with 0.1% SDS. Disintegrations per min were measured by liquid
scintillation counting.
qRT-PCR to evaluate metallothionein II mRNA
A metallothionein II fragment identical to clone 6a.2 except
that it contained a 63-bp deletion was subcloned into pCDNA.3
(Invitrogen). RNA was transcribed using the T7 Promoter
Large Scale Transcription Kit (Novagen, Milwaukee, WI). After
phenol-chloroform extraction and ethanol precipitation, the synthesized
RNA was quantitated and analyzed using UV spectrophotometry and gel
electrophoresis.
RT reactions were performed on 200 and 300 ng deoxyribonuclease SAOS-2 total RNA plus a known amount of metallothionein II standard RNA in a 20-µl reaction. The reaction contained 1 x PCR buffer (Life Technologies, Inc.), 3.75 mM MgCl2, 1.25 µM metallothionein II-specific reverse primer (5'-GGAATATAGCAAACGGTCAGGGTC-3'), 0.5 mM dNTPs, 1 mM dithiothreitol, 20 U RNasin (Promega Corp.), and 200 U Superscript II reverse transcriptase (Life Technologies, Inc.). Reactions were incubated at 42 C for 15 min, followed by 5 min at 99 C and 5 min on ice before amplification.
PCR was initiated by adding 80 µl of a master mix containing metallothionein II-specific forward primer (5'-GGCTCCTGCAAATGCAAAGAG-3') directly to the 20-µl RT reaction. The final concentrations of reagents in the 100-µl PCR reaction were 0.25 µM of each metallothionein II-specific primer, 1 x PCR buffer (Life Technologies, Inc.), 0.1 mM dNTPs, 1.5 mM MgCl2, and 0.5 U Taq DNA polymerase (Life Technologies, Inc.). Two-step PCR was carried out in a PE 9600 for 40 cycles as follows: 95 C for 30 sec and 64 C for 1.5 min. Samples were incubated at 64 C for 10 min after amplification.
PCR products were separated and analyzed on a reverse phase ion pair HPLC DNASep column (Sarasep, San Jose, CA). The elution system was a gradient of acetonitrile in 0.1 M triethylammonium acetate at a flow rate of 0.7 ml/min. The acetonitrile gradient increased from 14.6% to 16.6% over 5 min. The amount of product from standard and native RNAs was determined by UV absorbance detection at 254 nm, and signal was analyzed by an on-line integrator. From the chromatograms, the ratio of the area under each peak was used to determine the ratio of the amount of input metallothionein II standard RNA to the amount of native metallothionein II message in the SAOS-2 RNA.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, mRNA when assessed by RT-PCR
(Fig. 1
). As a positive control, GAPDH
mRNA was coamplified in these reactions. Whereas both cell lines
contained GAPDH mRNA, these two cell lines contained ERß and no
ER.
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).
Clone 6a was sequenced, and a nucleotide homology search revealed 98%
identity with human metallothionein II. Over the coding sequence,
however, there were no amino acid mismatches.
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). In the SAOS-2 cells, 10
nM 17ß-estradiol up-regulates metallothionein II mRNA as
much as 14-fold (Fig. 3). This up-regulation in SAOS-2 cells has been
repeated several times with a range of 3.5- to 14-fold and an
EC50 of 4.6 ± 2.7 nM, as
determined by qRT-PCR. If SAOS-2 cells are treated with 10
nM 17ß-estradiol, and RNA was prepared at different times
posttreatment, the first discernible increase in metallothionein II
message occurred after 8 h and was maximal by 24 h (Fig. 4).
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). Although comparable amounts of
receptor protein were expressed with and without cycloheximide
treatment, as measured by a whole cell binding assay (Fig. 5B),
estradiol did not increase metallothionein II mRNA expression (Fig. 5, C and D).
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. As illustrated in Fig. 6, endogenous levels of ERß or
overexpressed levels of ER were ineffective at mediating
17ß-estradiol regulation of metallothionein II. Whole cell ER binding
assay was used to verify expression of protein, and comparable results
were obtained for cells expressing ERß and ER (data not shown). As
an additional positive control for expression of functional ER
protein, SAOS-2 cells were coinfected with an Ad5 ERE-TK-luciferase
reporter gene and Ad5 ER, then treated with 17ß-estradiol with or
without ICI-182780. As shown in Fig. 7, 17ß-estradiol up-regulated reporter gene activity, and this
regulation was blocked by the antiestrogen ICI-182780. Therefore, the
inability of ER to regulate metallothionein II in SAOS-2
cells is not because the protein is improperly expressed. In LNCaPLN3
cells, ER did up-regulate metallothionein II mRNA, but to a lesser
extent than ERß (2- vs. 6-fold).
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). The antiprogestin/antiglucocorticoid,
RU486, did not block regulation by 17ß-estradiol (data not
shown).
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lists the cell lines tested. All cells
lines were infected with Ad5 hERßL to
overexpress ERß and were treated with 10 nM
17ß-estradiol. Receptor expression was monitored for most cell lines
by Northern blot or RT-PCR to ensure successful infection. Although
cells had varying levels of basal metallothionein II expression, the
response has only been detected in SAOS-2 and LNCapLN3 cells to
date.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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. The
reverse situation has recently been described, where ER, but not
ERß, can direct osteopontin from an SF-1 response element
(31). Interestingly, ER does not regulate
metallothionein II mRNA in SaOS-2 cells, but does so in LNCaPLN3 cells.
Clearly there are differences between these cell lines, perhaps in
their expression of coregulatory proteins, that determine ER’s
ability to direct metallothionein II transcription. Metallothioneins are low mol wt, cysteine-rich proteins that bind metals such as cadmium, copper, and zinc. Although the first metallothionein was discovered over 40 yr ago (32), debate continues as to their function. Several proposals have been made, and these include protection from metal toxicity, regulation of zinc and copper homeostasis, and defense against oxidative stress. Regulation of energy balance has also been implicated, because after reaching sexual maturity, metallothionein I and II knockout mice become obese (33).
Recently, studies have detailed how metallothionein may act to regulate zinc homeostasis in cells. Using purified zinc-dependent enzymes such as Escherichia coli alkaline phosphatase, bovine carboxypeptidase A, and sheep sorbitol dehydrogenase, two recent publications show how agents in the cellular milieu can facilitate exchange between zinc complexed with metallothionein and (metal-depleted) apo-enzymes (34, 35). Citrate and glutathione can influence the direction of zinc transfer and thus regulate enzyme activity depending on the redox state of the cell. Although not an enzyme, the ER also requires zinc for activity, and ER from MCF-7 cells can reversibly exchange zinc with purified metallothionein (36).
A myriad of agents can regulate metallothionein levels, including
glucocorticoids and metals (for review, see Ref. 37).
Estrogens are not classical regulators of metallothionein, but several
reports have appeared in the literature associating them. Estrogens are
reported to slightly antagonize dexa-methasone’s up-regulation of
metallothionein in cultured bovine pulmonary artery endothelial cells
(38). In the arctic char, treatment with 17ß-estradiol
did not affect basal metallothionein levels in several target organs
(39), whereas in the rainbow trout, it caused a
redistribution of cadmium from the bone and liver to the gut, gill, and
muscle (40). In rats, a 2-week treatment of female rats
with 17ß-estradiol up-regulated a copper-binding protein in
intestinal mucosa that reduced the amount of copper absorbed into the
plasma. As the mol wt of this protein was about 10K, the researchers
suggested that it may be metallothionein (41). Finally,
metallothionein was identified in a subtractive hybridization screen of
uterine mRNAs regulated after a single injection of ethinyl
estradiol (42). Although the isoform is not
identified, a metallothionein transcript increased 3-fold between 4–8
h after estrogen stimulation. In addition, it is not clear which
receptor type effects this regulation, because ER, not ERß, is the
most abundant ER in the rat uterus (1, 13).
As the functions of metallothionein and ERß are only beginning to be elucidated, understanding the significance of their association is difficult at this time. Future studies, such as additional gene profiling of tissues from estrogen-treated animals, may reveal that this regulation is not restricted to certain cell lines. In addition, as we learn more about the function of these proteins, a reason for estrogenic regulation of metallothionein may become apparent.
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Acknowledgments |
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Received June 6, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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. Mol Endocrinol 11:1486–1496 (ER-) and ß
(ER-ß) mRNA in the rat pituitary, gonad, and reproductive tract.
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rat central nervous system. J Comp Neurol 388:507–525[CrossRef][Medline]
via induction of Rab11 in uterine glands during
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ribonucleic acid in the wild type and ER-knockout mouse 1997.
Endocrinology 138:4613–4621 and ERß mRNA abundance in rats and the effect of
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.
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-knockout
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