From the Institute of Biochemistry and
§ Institute of Anatomy and Cell Biology, Humboldt
University Medical School Charité, Monbijoustraße 2, 10117 Berlin, Germany
Received for publication, September 5, 2002, and in revised form, October 28, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
A sperm nucleus glutathione
peroxidase (snGPx), which is closely related to the phospholipid
hydroperoxide glutathione peroxidase (phGPx), was recently discovered
in late spermatids. Both GPx isoforms originate from a joint ph/snGPx
gene, but their N-terminal peptides are encoded by alternative first
exons. The expression of the two enzymes is differentially regulated in
various cells, but little is known about the regulatory mechanisms. To
explore the tissue-specific regulation of expression of the two
isoenzymes, we first investigated their tissue distribution. Whereas
phGPx is expressed at low levels in many organs, snGPx was only
detected in testis, kidney, and in the human embryonic kidney cell line HEK293. Subcellular fractionation studies and immunoelectron microscopy revealed a cytosolic localization. To explore the mechanistic reasons
for the differential expression pattern, we first tested the activity
of the putative phGPx and snGPx promoters. The 5'-flanking region of
the joint ph/snGPx gene exhibits strong promoter activity. In contrast,
the putative snGPx promoter, which comprises 334 bp of intronic
sequences, lacks major promoter activity. However, it strongly
suppresses the activity of the ph/snGPx promoter. These data suggest
negative regulatory elements in the first intron of the ph/snGPx gene,
and DNase protection assays revealed the existence of several
protein-binding sites. The corresponding trans-regulatory
proteins (SP1, ERG1, GATA1, SREBP1, USF1, and CREBP1) were identified,
and in vivo binding of EGR1 and SREBP1 was shown by
chromatin immunoprecipitation. These data indicate for the first time
somatic expression of the snGPx and provide evidence for the existence
of intronic negative cis-regulatory elements in the
ph/snGPx gene. Our failure to detect an alternative snGPx promoter
suggests that transcription of the ph/snGPx gene may be regulated by a
joint basic promoter. The decision, which GPx isoform is expressed in a
given cell, appears to be made by alternative splicing of a joint
primary transcript.
Selenium-containing glutathione peroxidases
(GPx)1 constitute a family of
antioxidative enzymes that are capable of reducing organic and
inorganic hydroperoxides to the corresponding hydroxy compounds
utilizing glutathione or other hydrogen donors as reducing equivalents
(1, 2). Up to now, five selenium GPx subtypes have been identified (3,
4), but there are only 4 distinct seleno GPx genes. The most recently
discovered member (5) of this enzyme family is the sperm nucleus
glutathione peroxidase (snGPx). It constitutes a 34-kDa selenoprotein
that is expressed at a high level in late spermatids of various species
and has been implicated in chromatin condensation during sperm
development and in antioxidative protection of sperm DNA. It is
localized in the spermatid nuclei, and its expression is strongly
impaired under selenium deficiency (5, 6). The snGPx shares a high degree of protein-chemical similarity (5, 7) with the previously described phospholipid hydroperoxide glutathione peroxidase (phGPx), because both GPx isoforms originate from a joint ph/snGPx gene (5). In
fact, their C-terminal peptides (about 60% of the snGPx) are identical
and they are encoded for by exons 2 to 7 of the joint ph/snGPx gene. In
contrast, the N-terminal peptides of the two enzymes originate from
alternative first exons (Scheme 1). The
N-terminal peptide of snGPx 7 (15 kDa) contains an arginine-rich nuclear insertion sequence, which enables nuclear localization and is unique among selenoproteins (5). In contrast, the N-terminal peptide of the phGPx is much shorter (about 3 kDa) and lacks a nuclear
import sequence. Because both GPx isoforms are encoded for by the same
gene, the two enzymes may be considered splicing variants of a joint
ph/snGPx pre-mRNA (3, 4). However, it is not clear whether
expression of the two proteins really involves the formation of a joint
pre-mRNA or whether there are distinct transcription initiation
sites and alternative promoters. Considering the given genetic
composition (Scheme 1) it may be speculated that the 5'-flanking region
of the ph/snGPx gene is of functional relevance only for phGPx
transcription. In contrast, the immediate 5'-flanking region of exon
E1b (first snGPx exon) that includes the 5' part of intron 1 (I1a in
Scheme 1) and exon 1a (first phGPx exon) may contain regulatory
elements for snGPx transcription. In fact, stringent in
silico structural analysis of I1a indicated the presence of
putative transcription factor binding sites. These data suggested the
existence of an independent transcriptional regulation of phGPx and
snGPx expression.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (11K):
[in a new window]
Scheme 1.
Structure of the ph/snGPx gene and putative
transcription factor binding sites in intron I1a. The gray
boxes represent the phGPx exons (E1 to E7) and the introns (I1 to
I6) are indicated by the line between the exons. The start
codons for the mitochondrial and cytosolic phGPx isoforms are shown as
5'-ATG and 3'-ATG, respectively. ATG(sn)
represents the translational initiation site of the snGPx.
Sec indicates the localization of the catalytic
selenocystein and secis marks the selenocystein insertion
sequence required to prevent premature termination of translation at
the amber triplet encoding for the selenocystein. Analysis of the
transcription factor binding sites was carried out with the
MatInspector program.
The various isoforms of seleno-GPx show a remarkable tissue-specific expression pattern (3). The phGPx is expressed at relatively low levels in most cells and tissues tested (8, 9) but is much more prominent in testis (10). By contrast, snGPx mRNA has only been detected in testis and its expression appears to be restricted to late stages of spermatogenesis (5). Unfortunately, the regulatory mechanisms involved in high level expression of phGPx in germinative cells and repression of snGPx expression in most somatic tissues are unknown. Moreover, the molecular processes of transcriptional activation of snGPx expression during late sperm development remain to be investigated.
The lack of experimental data on the regulation of expression of the
ph/snGPx gene and the remarkable tissue-specific expression pattern of
the two GPx isoforms prompted us to study the regulatory mechanisms
involved in ph/snGPx expression. Here we report for the first time that
snGPx does not only occur in the nucleus of late spermatids but also in
the cytosol of interstitial kidney cells. Moreover, we found that the
regulation of expression of the joint ph/snGPx gene appears to be a
complex process that requires cis-regulatory elements in the
5'-flanking region of the ph/snGPx gene and the binding of negative
trans-regulatory proteins, such as SREBP1 and EGR1, to
intronic sequences (I1a). The lack of major promoter activity in the
5'-flanking region of the first snGPx exon argues against the existence
of alternative phGPx/snGPx promoters but strongly suggests alternative
splicing of a joint ph/snGPx pre-mRNA as mechanistic reason for
differential expression of the two enzymes.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials-- The chemicals used were from the following sources: trisodium citrate dihydrate, magnesium sulfate heptahydrate, Triton X-100, bovine serum albumin, penicillin-streptomycin solution, and fetal bovine serum from Sigma (Deisenhofen, Germany); sodium hydroxide, sodium chloride, and Tris from Merck (Darmstadt, Germany); PWO DNA polymerase, agarose, ampicillin, avian myeloblastosis virus reverse transcriptase, and proteinase inhibitor mixture tablets (complete, Mini) from Roche Molecular Diagnostics (Mannheim, Germany); restriction endonucleases and the DNA molecular weight markers (100 bp and 1 kb) from New England Biolabs GmbH (Schwalbach, Germany); Servalyt 3-10, 6-8, and 2-4 from Serva (Heidelberg, Germany); PanScript DNA polymerase from PAN BIOTECH GmbH (Aidenbach, Germany); Advantage 2 polymerase from Clontech (Palo Alto, CA); Bacto-yeast extract, Bacto-agar, and Bacto-tryptone from Difco (Detroit, MI); rainbow molecular weight markers and Hybond-N blotting membrane from Amersham Biosciences (Freiburg, Germany); and Dulbecco's modified Eagle's medium from Invitrogen (Karlsruhe, Germany). The primers were custom synthesized by TIB MOLBIOL (Berlin, Germany).
Cell Lines and Culture Conditions-- Human embryonic kidney cells 293 (HEK293) were obtained from the German Collection of Microorganisms and Cell Cultures (Braunschweig, Germany). The cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% (v/v) fetal calf serum and antibiotics (100 units/ml penicillin and 100 µg/ml streptomycin) at 37 °C under 5% CO2.
Animal Experiments--
Mice (C57BL/N6, 10-15 weeks old) were
sacrificed by diethyl ether inhalation and the major organs were
prepared. After extensive washing with PBS to remove blood, the tissue
was homogenized on ice in 5 volumes of TBS (25 mM Tris-HCl,
130 mM NaCl, 5 mM KCl, protease inhibitor
mixture, pH 7.4) in a glass Dounce homogenizer. Cell debris was spun
down at 20,000 × g and aliquots of the supernatants were subjected to immunoblotting. For nuclear extracts HEK293 cells,
mouse testis or kidney were lysed in 10 mM HEPES buffer, pH
7.9, containing 10 mM KCl, 1.5 mM
MgCl2, 0.1 mM EDTA, 0.35 M sucrose,
0.5 mM DTT, and a proteinase inhibitor mixture (see "Materials"). After 10 min on ice the cells were homogenized in a
glass Dounce homogenizer and recentrifuged at 7000 rpm. The nuclear
pellet was resuspended in 20 mM Hepes buffer, pH 7.9, containing 10% glycerol, 420 mM NaCl, 1.5 mM
MgCl2, 0.1 mM EDTA, 0.5 mM DTT, and
proteinase inhibitor mixture and was incubated on ice for 30 min. The
suspension was centrifuged at 10,000 × g for 15 min at
4 °C and the supernatant was stored at 80 °C. The protein
concentration in all samples was measured by colorimetric assay
(Roti-Quant, Roth, Karlsruhe, Germany).
Reverse Transcriptase-Polymerase Chain Reaction-- Total RNA was isolated from mouse tissues or HEK293 cells using the RNA MIDI or MINI Kits from Qiagen (Hilden, Germany) and 3 µg were reverse-transcribed at 37 °C for 90 min in 45 µl of Tris-HCl buffer, pH 8.2, containing 8 mM MgCl2, 30 mM KCl, 1 mM dithiothreitol, 100 µg/ml bovine serum albumin, 30 units of RNase inhibitor, 0.166 mM dNTPs, 150 pmol of oligo(dT) primer, and 15 units of reverse transcriptase. To stop the reaction samples were heated to 95 °C for 10 min. RT-PCR signals were normalized for the signals of the human or murine glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which were amplified with the following primer combinations: human GAPDH, 5'-TCGGAGTCAACGGATTTGGTCGTA-3' and 5'-ATGGACTGTGGTCATGAGTCCTTC-3'; murine GAPDH, 5'-TCGGTGTGAACGGATTTGGCCGTA-3' and 5'-ATGGACTGTGGTCATGAGCCCTTC-3'. The following primer combinations were used for amplification of the murine and human snGPx: 5'-TCGCCGGATGGAGCCCATTCCT-3' and 5'-ACGCAGCCGTTCTTATCAATGAGAA-3' (murine); 5'-CCGGCGGAAGAAGCCCTGTCC-3' and 5'-CGAATTTGACGTTGTAGCCCGCG-3' (human). For amplification, 1 µl of the reverse transcription reaction was used. After initial denaturation for 4 min at 94 °C, 30 cycles of PCR were performed. Each cycle consisted of a denaturing period (40 s at 94 °C), an annealing phase (murine GAPDH: 60s at 69 °C, human GAPDH: 60s at 66 °C, snGPx: 60 s at 67 °C), and an extension period (120 s at 72 °C). After the last cycle, all samples were incubated for an additional 10 min at 72 °C. PCR products were separated by 2% agarose gel electrophoresis, and the DNA bands were stained with ethidium bromide.
Two-dimensional Electrophoresis and Immunoblotting-- The cytosolic proteins of kidney homogenates were mixed with 10 µl of sample buffer (1.43 g/ml urea, 10 mg/ml dithiothreitol, 10% (w/v) Triton X-100, 107 µl/ml Servalyt 3-10, 26 µl/ml Servalyt 6-8, 16 µl/ml Servalyt 2-4) and then subjected to isoelectric focusing (first dimension) in acrylamide cylinders, in which a linear pH gradient between 2 and 10 was adjusted. SDS-PAGE (second dimension) was subsequently carried out in a 12.5% polyacrylamide gel. After SDS-PAGE, the two-dimensional chromatograms were either stained with 0.2% (w/v) Coomassie Brilliant Blue R-250 or were transferred to a nitrocellulose membrane NC45 (Serva, Heidelberg, Germany) by a semidry blotting procedure. To stain the immunoblots the nitrocellulose membrane was first incubated for 1 h with a 1:4000 dilution of a monoclonal antibody raised against the Sec46Cys mutant of the human cytosolic phGPx. This antibody prepared in our laboratory strongly cross-reacts with the rat and murine phGPx but does not distinguish between phGPx and snGPx. After extensive washing with PBS, the membrane was incubated with a peroxidase-labeled goat anti-mouse IgG antibody (Sigma) and immunoreactive protein bands were visualized with a 3,3'-diaminobenzidine/H2O2 mixture.
Functional Promoter Assays--
The pBlueTOPO reporter vector
(Invitrogen, Groningen, The Netherlands) containing the
-galctosidase as reporter gene was used for analysis of promoter
activity of subcloned promoter fragments. For the assays, putative
promoter fragments of phGPx and snGPx were amplified by PCR and
subcloned into the pBlue-TOPO reporter vector following the user's
manual. PCR was carried out with a Biometra TRIO Thermoblock 2.51BB
(Biometra, Göttingen, Germany) using purified plasmid DNA
(Qiagen, Hilden, Germany) of the subcloned ph/snGPx gene as template
and the following primer combinations were used (Fig. 5): 5'-flanking
region of the ph/snGPx gene (construct C1, 325 bp),
5'-CGCGTCCCTATCACTGGGGCATG-3' and 5'-TGGTGCCTGCCAGACCAGGCG-3'; immediate 5'-flanking region of the first snGPx exon (construct C2, 334 bp), 5'-TGGGCTACTGGGAACTTGGAGGA-3' and 5'-ATGCCCGCCGGTCTGTGCGTC-3'; extended 5'-flanking region of the first snGPx exon (construct C3, 662 bp) 5'-CGCGTCCCTATCACTGGGGCATG-3' (same as for construct C1) and
5'-ATGCCCGCCGGTCTGTGCGTC-3'. For transient transfection HEK293 cells
were resuspended at a density of 0.5 × 106 cells/5 ml
of medium and plated in 60-mm culture dishes. After a culturing period
of 24 h cells were transfected with a calcium phosphate
transfection kit (Invitrogen). For transient transfection we used 10 µg of the
-galactosidase promoter construct and 5 µg of an
internal reference plasmid containing the luciferase cDNA (pGL3,
Promega, Mannheim, Germany). After 72 h, cells were harvested and
both
-galactosidase and luciferase activities were assayed. The
-galactosidase activities were normalized for transfection efficiency using the data of the luciferase assay.
DNase I Protection Assay--
A genomic fragment containing
intron I1 of the ph/snGPx gene was subcloned into the NcoI
and PstI site of the Litmus29 vector (Promega). After
excision with XbaI and BamHI the probe was
radioactively labeled using T4 polynucleotide kinase. The labeled probe
was then treated with PstI or NcoI to achieve
selective labeling of either the sense or antisense strand. Aliquots of
the labeled probe containing 10,000-20,000 cpm of
32P-labeled DNA were added to reaction mixtures consisting
of a 40 mM Hepes buffer, pH 7.9, containing 100 mM KCl, 12.5 mM MgCl2, 1 mM EDTA, 20% glycerol, 1 mM DTT, 1 µg of
poly(dI-dC), and 10-20 µg of nuclear proteins in a total volume of
50 µl. The mixture was kept on ice for 30 min before addition of 50 µl of 5 mM CaCl2, 10 mM
MgCl2, and 2-5 units of DNase I depending on the input of nuclear proteins. Digestion was carried out at 4 °C for 5 min and
terminated by adding 100 µl of a solution containing 200 mM NaCl, 20 mM EDTA, and 1% SDS. DNA was
phenol extracted and precipitated with ethanol. The digestion products
were analyzed on 6% acrylamide sequencing gels containing 8 M urea. The protected sequences were identified by
comparison to the migration of Maxam-Gilbert sequencing products. Gels
were dried and exposed to x-ray films overnight at 80 °C.
Electrophoretic Mobility Shift Assay (EMSA) and Supershifts-- EMSA was carried out with the DIG Gel Shift Kit (Roche Molecular Diagnostics) following the users manual. In the first step single-stranded complementary oligonucleotides containing the binding sequences for transcription factors were annealed and end-labeled with digoxigenin. The labeled probes (48 fmol of double-stranded oligonucleotides) were then incubated for 30 min at 4 °C with 10 µg of nuclear extract proteins in 40 mM Hepes buffer, pH 7.9, containing 100 mM KCl, 12.5 mM MgCl2, 1 mM EDTA, 20% glycerol, 1 mM DTT, 1-2 µg of poly(dI-dC), 0.1-0.2 µg of poly(L) lysine (total assay volume of 15 µl). Then the mixtures were subjected to electrophoresis on a 6% polyacrylamide gel with 0.5-fold TBE running buffer. The digoxigenin-oligonucleotide/protein complexes were transferred to a Hybond-N blotting membrane (Amersham Biosciences) and the shift bands were visualized following the users manual. For competition studies unlabeled probe competitor or consensus oligonucleotides (Santa Cruz Biotechnology, Santa Cruz, CA) were added at a 166-fold excess over the digoxigenin-labeled probe. For supershift studies, polyclonal antibodies from Santa Cruz Biotechnology were used. When either competitor or antibodies were used, they were preincubated with the nuclear extracts at 4 °C for 10 or 60 min before the addition of the digoxigenin-labeled probe DNA.
Chromatin Immunoprecipitation--
107 HEK293 cells
were washed twice with PBS and the cellular proteins were cross-linked
to DNA by adding formaldehyde to a final concentration of 1% for 15 min at room temperature. Cells were subsequently washed, removed from
the culture dishes by scraping, and resuspended in 1 ml of PBS. Cells
were lysed in 1.5 ml of lysis buffer (10 mM Hepes buffer,
pH 7.9, containing 10 mM KCl, 1.5 mM
MgCl2, 0.1 mM EDTA, 0.5 mM
DTT) and the proteinase inhibitor mixture. After 10 min on ice
the cells were homogenized in a glass Dounce homogenizer and
centrifuged at 7,000 rpm. The nuclear pellet was resuspended in 20 mM Hepes buffer, pH 7.9, containing 10% glycerol, 420 mM NaCl, 1.5 mM MgCl2, 0.1 mM EDTA, 0.5 mM DTT, and proteinase inhibitor
mixture and was sonicated on ice to fragment the genomic DNA. The
mixture was then centrifuged to remove cell debris and aliquots were
stored at 80 °C. For chomatin immunoprecipitation aliquots were
diluted in RIPA buffer (50 mM Tris-HCl buffer, pH 7.5, containing 150 mM NaCl, 0.1% sodium desoxycholate, 1%
Triton X-100, 0.25 mM EDTA, and the proteinase inhibitor
mixture) and samples were incubated overnight at 4 °C with 2.5 µl
of a rabbit polyclonal anti-Egr1 antibody (588) or with a rabbit
anti-SREBP1 antibody (H-160) obtained from Santa Cruz Biotechnology.
Immune complexes were precipitated by adding 50 µl of protein
A-agarose (Santa Cruz Biotechnology) for 3 h with low speed
spinning. Precipitates were washed twice with 1 ml of RIPA buffer,
twice with 1 ml of PBS, and DNA-protein complexes were eluted from the
agarose by a 15-min incubation with 250 µl of elution buffer (1%
SDS, 100 mM NaHCO3). Elution was repeated, the
eluates were pooled, NaCl was added to reach a final concentration of
0.3 M, and the samples were incubated at 65 °C for
4 h to reverse formaldehyde-induced cross-linking. The proteins
were then digested with 2 µl of proteinase K (20 mg/ml) in 40 mM Tris-HCl buffer, pH 7.5, containing 10 mM EDTA. The samples were incubated for 1 h at 60 °C and the DNA was extracted with phenol/chloroform followed by ethanol precipitation. The DNA pellet was resuspended in 50 µl of sterile H2O
and 10 µl of DNA solution was used as a template for PCR. The
following primer combinations were designed to amplify the precipitated binding regions of the transcription factors (Fig. 11) and 35 PCR cycles were run: P1 (5'-CCATGGTGGGCTACTGGGAACTT-3'), P1'
(5'-TCCCGCGCCGAGGCCTAGCC-3'), P2 (5'-AGGCCTCGGCGCGGGAGGTC-3'), P2'
(5'-ATGCCCGCCGGTCTGTGCGTC-3'). The PCR products were separated on
2% agarose gel and visualized by ethidium bromide staining.
Immunohistochemistry-- For immunohistochemistry the animals (two rats and two mice) were perfused with ice-cold 4% (w/v) paraformaldehyde (dissolved in PBS). Kidneys were prepared, cryopreserved in 20% sucrose for 24 h at 4 °C, and frozen in the gaseous phase of liquid nitrogen. Cryostat sections of 10-20 µm thickness were prepared and quenched in 50 mM NH4Cl/PBS for 30 min. The sections were blocked with 10% goat serum, 0.1% saponin in PBS for 1 h at room temperature and exposed to the primary monoclonal anti-PH-GPx antibody (1:200 diluted) overnight at 4 °C. After washing in 0.1% saponin/PBS, sections were incubated overnight at +4 °C with a biotinylated anti-mouse IgG antiserum (Vector Laboratories, Burlingame, CA; diluted 1:250). For staining, the sections were exposed to avidin-biotin peroxidase complex reagent (Vector Laboratories) for 2 h at room temperature, and the immunoreaction was visualized with 3,3'-diaminobenzidine as a chromogen (incubation for 8 min at room temperature with 0.07% 3,3'-diaminobenzidine and 0.001% hydrogen dissolved in 0.1 M PBS). Prior to inspection the sections were counterstained with hematoxylin eosin. Control staining was performed appropriately using a diluted preimmune serum.
Immunoelectron Microscopy-- For ultrastructural studies, ph/snGPx-immunostained sections were osmicated (1% OsO4 in 6.84% sucrose dissolved in PBS) for 5 min, dehydrated in graded ethanol, and flat embedded in Epon medium (Merck, Darmstadt, Germany) between silane-coated slides. Ultrathin sections were cut on a ultratome, mounted on single slot grids coated with a Formvar film, and stained with lead citrate and uranyl acetate. A Zeiss EM 900 electron microscope (Zeiss, Jena, Germany) was used for examination.
Activity Assay--
The phospholipid hydroperoxidase activity
was assayed measuring the decrease in absorbance at 340 nm with a
coupled optical test (11). The assay mixture consisted of 0.1 M Tris-HCl, pH 7.6, containing 5 mM EDTA, 0.1%
Triton X-100, 0.2 mM NADPH, 3 mM glutathione,
and 1 unit of glutathione reductase in a total assay volume of 0.2 ml.
The enzyme preparation (5-20 µl) was preincubated at room
temperature in the assay buffer for 10 min and then the reaction was
started by addition of 10 µl of a methanolic solution of high
performance liquid chromatography-purified phospholipid hydroperoxide
reaching a final substrate concentration of 50 µM. The
hydroperoxy phospholipids were prepared by incubating for 10 min
soybean phospholipids in 0.2 M borate buffer, pH 9, containing 3 mM sodium desoxycholate and 1 mg/ml soybean
lipooxygenase. After lipid extraction the conjugated dienes were
prepared by reverse phase-high performance liquid chromatography (12)
and the salt included in the solvent system was removed by solid-phase
extraction. Final quantification of the peroxide preparation was
carried out spectrophotometrically measuring the absorbance at 235 nm
using a molar extinction coefficient of 25,000 (M
cm)1.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Tissue-specific Expression of snGPx and Its Intracellular
Localization--
It has been previously reported that phGPx is
ubiquitously expressed in small amounts in many cells and tissues (3,
8, 9) but at much higher levels in testis (10, 13). In addition, Northern blot analysis suggested a high level expression of the snGPx
mRNA in testis and its absence in liver, kidney, spleen, brain, and
other organs (5). Within the testis the enzyme was mapped predominantly
to the nuclei of late spermatids (5). Because spermatids are not
suitable for mechanistic studies, the tissue-specific expression
pattern of the snGPx mRNA was reinvestigated by semiquantitative
RT-PCR. Here we found that the snGPx mRNA is present at a high
level in testis and at lower levels in kidney (Fig.
1). In all other major organs (ileum,
skin, heart, brain, liver, lung, stomach, spleen, muscle, and
submandibular gland) we were unable to detect this messenger (not
shown). In some experiments faint snGPx bands were observed in lung and
stomach. In the snGPx-positive tissues (testis and kidney) we
consistently detected an additional PCR product migrating with a higher
molecular weight (Fig. 1). This PCR fragment was subcloned and
sequenced, and the data indicated an incomplete splicing product that
still contained intron I1b (Scheme 1). However, all other introns
(I2-I6) had been spliced out completely. These data suggest problems
of the splicing machinery during snGPx expression. It is likely that
proteins binding to I1a of the joint ph/snGPx pre-mRNA may prevent
proper splicing of the primary transcript.
|
For mechanistic studies on the regulation of gene expression, solid tissues have several limitations. Thus, we searched for a permanent cell line, in which the snGPx is expressed. For this purpose several human and murine cell lines were tested and expression of the snGPx mRNA was found in the human embryonic kidney cell line HEK293 (data not shown). Interestingly, this cell line also expresses phGPx as indicated by RT-PCR with phGPx-specific primer combinations.
To confirm the tissue-specific expression of snGPx on the protein
level, immunoblots with a monoclonal anti-ph/snGPx antibody were
carried out. For this purpose 20,000 × g supernatants
were prepared from murine tissues and equal protein amounts were
applied to SDS-PAGE. From Fig.
2A it can be seen that
immunoreactive proteins migrating in the expected 34-kDa region were
present in the kidney lysis supernatant suggesting snGPx expression in
this organ. In a separate experiment electrophoretic co-migration of
the kidney snGPx with an snGPx standard prepared from human spermatids
was observed. Moreover, an immunoreactive band of minor intensity was
detected in the 20-kDa region that may be because of phGPx. By
comparison, large amounts of this low molecular weight protein were
found in the testis high speed supernatants. These data indicate that
in testis cytosol the phGPx predominates and the snGPx appears to be
absent.
|
The kidney snGPx exhibited a catalytic activity for reduction of hydroperoxy phosphatidylcholine of 0.057 units/mg of protein. This specific activity is similar to previously reported values (0.028 units/mg) (11). Our attempts to purify the native snGPx from murine kidney supernatants by conventional protein purification techniques were not successful. Running the supernatant over sequential Mono-Q FPLC and gel filtration resulted only in 8-fold enrichment of the immunoreactive material with substantial loss of enzymatic activity. In the most active FPLC fractions the specific activity could only be enriched by a factor of 2 (0.12 units/mg) compared with the lysis supernatant (see above).
It was previously shown that phGPx forms covalently linked polymers (10) that may migrate at a higher molecular weight region. Thus, the 34-kDa band observed in kidney cytosol may be because of a phGPx dimer instead of a snGPx monomer. To distinguish between these two protein species we carried out two-dimensional electrophoresis (Fig. 2B) and found that the immunoreactive material from the kidney lysate supernatant migrated with an apparent isoelectric point of about 7. However, a more alkaline isoelectric point has been reported for phGPx (7, 10). It should be stressed that in two-dimensional electrophoresis, we did not observe major expression of the 20-kDa phGPx. Comparison of Coomassie-stained immunoblots revealed that snGPx is not a major protein in kidney cytosol. In the region where the immunoreactive material was detected, significant protein was not observed.
The relatively large amounts of snGPx detected in kidney cytosol were somewhat surprising because the enzyme contains a nuclear import sequence and thus, should be localized preferentially in the nuclei. However, subcellular fractionation studies combined with Western blot analysis indicated the predominant cytosolic localization of the snGPx in kidney and its lack in the nucleus (Fig. 2C).
The available ph/snGPx antibodies do not distinguish between the two
isoforms and thus, it was not possible to discriminate between the two
GPx isoforms using immunohistochemistry. Fortunately, the Western blots
indicated predominant expression of the 35-kDa snGPx in the kidney and
thus, immunohistochemistry of kidney cross-sections will mainly reflect
the cellular and subcellular distribution of the snGPx. In murine
kidney, ph/snGPx is mainly expressed in cortical and medullary
interstitial cells (Fig. 3). In contrast, the glomeruli and the epithelial cells of the tubuli and collecting ducts were free of immunoreactive material. Similar results were also
obtained with rat kidneys (data not shown). These data contrast with
previous reports suggesting the expression of the phGPx in renal
epithelial cells (14). Because of the higher density of interstitial
cells in the medulla the level of ph/snGPx expression is apparently
higher in this renal compartment (panel A). In additional experiments we investigated the intracellular distribution of the snGPx
in renal cortex and medulla by immunoelectron microscopy. From Fig.
4 the immunoprecipitate is seen to
localize in the cytoplasm and below the cellular membrane of
interstitial (panel A) and vascular endothelial cells
(panel C). Here again, the tubular epithelium did not show
any immunoreactivity (panel B).
|
|
Functional Promoter Assays--
If differential expression of
phGPx and snGPx in a given cell is transcriptionally regulated,
distinct transcription initiation sites and different basic promoter
regions for the two isoenzymes would be expected. To find out whether
there is significant promoter activity in the 5'-flanking region of
exon 1b (Scheme 1) functional promoter assays were carried out. For
this purpose -galactosidase-based reporter gene constructs were
transfected into HEK293 cells and
-galactosidase activity was
assayed (Fig. 5). For control studies the
-galactosidase reporter gene construct without inserted
oligonucleotide was used (construct C0). Construct C2, which contained
a 334-bp fragment (I1a) of the 5'-flanking region of E1b (putative
minimal snGPx promoter), only exhibited minor promoter activity. When compared with the putative ph/snGPx promoter, which involves the 5'-flanking region of the ph/snGPx gene and the 5'-untranslated region
of the corresponding mRNA (construct C1), its activity was lower by
2 orders of magnitude. In additional experiments the impact of I1a on
the activity of the phGPx promoter was tested (construct C3).
Surprisingly, I1a was found to suppress the activity of the phGPx
promoter in HEK293 cells by more than 95%. These data indicate that
I1a may contain negative regulatory elements for ph/snGPx expression
and that the corresponding trans-acting inhibitor proteins
may be present in HEK293 cells.
|
Identification of Protein Binding Sites in I1a--
The results of
the functional promoter assays suggests that negative regulatory
proteins bind to I1a. To investigate DNA/protein interactions in this
region we carried out DNase protection assays using nuclear extracts of
mouse testis and HEK293 cells as sources for the
trans-regulatory proteins. The data presented in Fig. 6 indicate: (i) the existence of at least
five distinct protein-binding regions (FP1-FP5) in I1b and (ii) the
presence of the corresponding trans-acting proteins in the
nuclei of testis and HEK293 cells. The lack of DNase protection by
albumin (data not shown) indicates the specificity of protein
binding.
|
Next, the sequences of the different protein-binding sites were
determined and examined for the presence of transcription factor
binding motifs. As evident from Fig.
7A FP1 contains the consensus
binding sequences for the stimulating proteins (SP1-like factors) and
for the early growth response gene product/Wilms tumor suppressor gene
product (EGR/WT). EMSA were subsequently carried out to test the
presence of the corresponding trans-acting binding factors
in testis and HEK293 cells. With testis extracts we observed a single
high molecular weight shift band that was competed off by nonlabeled
competitor (Fig. 7B). By contrast, two shift bands were
detected when nuclear extracts of HEK293 cells were used. Addition of a
consensus oligonucleotide for SP1 caused a disappearance of the upper
SP1 shift band (Fig. 7C). In a similar experiment the strong
upper SP1 shift band from HEK293 extracts also disappeared, whereas the
lower band of minor intensity remained unaltered. The lower shift band,
however, was eliminated with a consensus oligonucleotide for the EGR
transcription factor family (Fig. 7B). Because HEK293 cells
are embryonic kidney cells adult kidney cells were also examined for
expression of the corresponding transcription factors. As shown in Fig.
7D we observed two shift bands when murine kidney nuclear
extracts were added to the assays. Here again, the upper band
disappeared completely when a SP1 consensus oligonucleotide was added,
whereas the lower band was quenched off in the presence of EGR
competitor. In a separate experiment an EGR1 antibody was added to the
reaction mixture and in this sample the EGR/WT shift band was removed.
However, we were not able to detect a clear supershift signal. Taken
together, these data suggest that footprint 1 is caused by the binding
of a member of the SP1 transcription factor family (15) and/or EGR1
(16) to appropriate binding sites in I1a. The differential quenching behavior of testis and HEK293 nuclear extracts suggest that the testis
extracts may not contain sufficient amounts of EGR1.
|
When the nuclear extracts of HEK293 cells and murine kidney were used as source of nuclear proteins an additional very low shift band (Fig. 7, B and C) was observed and this band does not appear when testis nuclear extracts were used. Unfortunately, it has not yet been possible to identify the corresponding trans-acting protein.
FP2 (Fig. 8A) contains a GATA1
binding motif (17) and EMSA indicated a single shift band when the
nuclear extracts from testis and HEK293 cells were used as source of
the binding proteins (Fig. 8B). This band was competed off
by addition of unlabeled competitor (Fig. 8B) and by a GATA
consensus oligonucleotide (Fig. 8C). In FP3 (Fig.
9A) a SREBP1-binding site was
detected (18) and EMSA experiments indicated single shift bands when
the nuclear extracts of testis and 293 cells were incubated with the
corresponding DNA probe (Fig. 9B). Addition of unlabeled
competitor eliminated the shift bands. A similar result was obtained
when a SREBP1 consensus oligonucleotide was added (Fig. 9C).
FP4 and FP5 (Fig. 10) contain binding
sites for USF1 (Fig. 10A) and CREB (Fig. 10B),
respectively (19, 20). After addition of the nuclear extracts of testis and HEK293 cells to the corresponding DNA probes we detected strong shift bands (Fig. 10, C and D). Addition of
unlabeled competitor (data not shown) and consensus oligonucleotides
caused a disappearance of these signals. Taken together, the EMSA
experiments indicate that cis-regulatory elements for the
binding SP1, EGR1, GATA1, SREBP1, USF1, and CREBP are present in the
5'-flanking region of E1b (Scheme 2) and
that the corresponding trans-regulatory factors are
expressed in murine testis and HEK293 embryonic kidney cells.
|
|
|
|
In Vivo Binding of ERG1 and SREBP1 to the 5'-Flanking Region of E1b
(Putative snGPx Promoter)--
The EMSA experiments indicated that the
above mentioned transcription factors are present in the nuclei of
testis and HEK293 cells and that they bind in vitro to the
corresponding cis-regulatory sequences in the 5'-flanking
region of E1b. To determine whether they also bind in vivo,
chromatin immunoprecipitation was carried out. For this purpose
DNA-binding proteins of HEK293 cells were covalently linked to genomic
DNA by treatment of the cells with formaldehyde. The DNA-protein
complexes were then sheared by sonication, and specific protein-DNA
complexes were immunoprecipitated with antibodies against SREBP1 and
ERG1. Covalent linkage was reversed and the precipitated double
stranded DNA was amplified by PCR with binding site-specific primers.
From Fig. 11B it can be seen that PCR signals were obtained when the DNA/protein adducts were immunoprecipitated with antibodies against SREBP1 and ERG1 indicating the in vivo binding of the transcription factors. In
contrast, the corresponding controls, in which immunoprecipitation was
performed without specific antibodies, did not show any PCR signal.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
GPxs constitute a family of functionally related enzymes and the phGPx and snGPx are two members of this family. Whereas phGPx is expressed at low levels in many cells and tissues (3, 8), snGPx was only detected in late spermatids of several mammalian species (5). In these cells the enzyme was predominantly localized in the nucleus and has been implicated in chromatin structuring during sperm development and in antioxidative defense of the DNA (4-6). In this paper we report for the first time that snGPx is also expressed in selected somatic cells (interstitial kidney cells). Immunohistochemical staining, immunoelectron microscopy, and subcellular fractionation studies indicate that the enzyme is located predominantly in the cytosol. This localization was rather surprising because snGPx contains a nuclear import peptide that has been proposed to target the protein to the nucleus. Currently, it is unclear why the enzyme is transported into the nucleus in late spermatids and why it remains in the cytosol in interstitial kidney cells. It is possible that nuclear import requires additional factors, which may not be present at the required concentration in the kidney cells.
The kidney-specific function of the snGPx has not yet been investigated. Because the enzyme is mainly expressed in interstitial cells but not in the glomeruli, the tubulus epithelium, or the collecting ducts, it is unlikely to play a role in urine production. Alternatively, the renal snGPx may be involved in antioxidative defense as it has been suggested before for other GPx isoforms (21). It is well known that there is an active oxidative metabolism in renal cortex, which may be accompanied by the formation of reactive oxygen species (22, 23).
Compared with other selenoperoxidases, which are encoded for by separate genes, phGPx and snGPx originate from a joint ph/snGPx gene (4, 5). Nevertheless, both isoforms are differentially expressed in various cells and tissues, but virtually nothing is known about the regulatory mechanisms. Here we report that the minimal promoter of the phGPx (cytosolic isoform), which comprises the 5'-flanking region of the ph/snGPx gene and the 5'-untranslated region of its mRNA, exhibits major promoter activity. By contrast, the promoter activity of the 5'-flanking region of E1b (putative snGPx promoter) was almost negligible. These data are difficult to reconcile with the existence of alternative phGPx/snGPx promoters. They rather suggest that transcription of the ph/snGPx gene is driven by joint regulatory sequences located upstream of the 3'-ATG of the ph/snGPx gene. Interestingly, the joint ph/snGPx promoter can be strongly down-regulated by inhibitory proteins that bind to intronic regulatory sequences of I1a. Possible suppressor protein candidates include members of several transcription factor families (e.g. SP, EGR, GATA, SREBP, USF, and CREBP). Functional intronic cis-regulatory elements have been reported to be involved in the regulation of expression of several genes (24, 25), but the detailed mechanisms of action are unknown.
RT-PCR of snGPx transcripts indicated the presence of incomplete splicing products that selectively contain I1a (Fig. 1). Interestingly, we have never observed splicing problems with introns 2-5 of the ph/snGPx gene. Although the mechanistic reasons for this incomplete splicing have not been investigated in this study, the identified transcription factors may be involved. It has been reported before that transcription factors that prefer to bind to double stranded DNA are also capable of binding to single stranded RNA (26, 27). Thus, they may be of regulatory importance for the splicing process. Alternative exons are frequently flanked by suboptimal splicing sites, and an increasing body of experimental evidence has indicated that effective outsplicing of alternative exons requires the participation of splicing enhancers and/or silencers in addition to the constitutive spliceosomal machinery (28). The presence of such auxiliary proteins may help to define the exact splicing sites, and may also be important for maturation-dependent and tissue-specific alterations of the protein expression patterns. cis-Regulatory pre-mRNA sequences that bind the trans-acting splicing factors may reside in the alternative exon itself (29), in the flanking introns (30, 31), and even in an adjacent exon (32). Inclusion of an alternative exon in the final mRNA is a multistep process that involves timely differentiated removal of the two flanking introns. In fact, in several alternatively spliced pre-mRNA species it has been demonstrated that one flanking exon is removed before the other (33, 34). Although the detailed splicing mechanism of the ph/snGPx pre-mRNA was not the focus of this study, we have identified functional cis-regulatory sequences in the 5'-flanking intron of alternative exon E1b, which may be related to the splicing process. Additional stringent in silico search for similar cis-regulatory elements in E1b also suggested the existence of SP1/WT1-binding sites and preliminary DNase protection assays suggested their functionality (data not shown). Similarly, the 3'-flanking intron of E1b (I1b) also contains binding sites for several transcription factors (GABP1, OCT1P1, and GATA3), but their functionality has not been tested. Taken together, these data suggest the presence of cis-regulatory elements in the alternative exon E1b and in the flanking introns I1a and I1b. Such a constellation is frequently found in genes containing alternative exons, and the cis-regulatory sequences may serve as binding regions for splicing enhancers and/or silencers.
In the light of these findings the functional cis-regulatory
elements identified in I1a and the corresponding
trans-regulatory proteins may be of dual functional
importance: (i) they may act as negative regulators of transcriptional
initiation of the ph/snGPx gene (Fig. 5) and (ii) they may exhibit
auxiliary function for alternative splicing of the joint ph/snGPx
mRNA. Work is in progress in our laboratory to provide more
detailed information on either process.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. R. Schmitt for some methodological considerations and M. Petzold for excellent technical assistance. Moreover, we are indebted to V. O'Donnell for critical reading of the manuscript and D. Behne for providing a standard of snGPx.
![]() |
FOOTNOTES |
---|
* This work was supported by Deutsche Forschungsgemeinschaft Grant SSP1087.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.
¶ To whom correspondence should be addressed: Institut für Biochemie, Universitätsklinikum Charité, Humboldt Universität, Monbijoustr. 2, 10117 Berlin, Federal Republic of Germany. Tel.: 49-30-450-528040; Fax: 49-30-450-528905; E-mail: hartmut.kuehn@charite.de.
Published, JBC Papers in Press, November 8, 2002, DOI 10.1074/jbc.M209064200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: GPx, glutathione peroxidase; phGPx, phospholipid hydroperoxide glutathione peroxidase; snGPx, sperm nucleus glutathione peroxidase; PBS, phosphate-buffered saline; SP1, stimulating protein 1; EGR/WT, early growth factor/Wilms tumor suppressor; GATA, GATA box; SREBP, sterol regulatory element-binding protein; USF, upstream stimulating factor; CREBP, cAMP-responsive element binding protein; FP, footprint; EMSA, electrophoretic mobility shift assay; HEK, human embryonic kidney; RT, reverse transcriptase; DTT, dithiothreitol; GAPDH, glyceraldehyde-3-phosphate dehydrogenase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Flohé, L. (1989) in Glutathione: Chemical, Biochemical and Medical Aspects, Part A (Dolphin, D. , Poulson, R. , and Avramovic, O., eds) , pp. 643-731, John Wiley & Sons Inc., New York |
2. | Arthur, J. R. (2000) Cell. Mol. Life Sci. 57, 1825-1835[Medline] [Order article via Infotrieve] |
3. | Brigelius-Flohé, R. (1999) Free Radical Biol. Med. 27, 951-965[CrossRef][Medline] [Order article via Infotrieve] |
4. | Kuhn, H., and Borchert, A. (2002) Free Radical Biol. Med. 33, 154-172[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Pfeifer, H.,
Conrad, M.,
Roethlein, D.,
Kyriakopouluos, A.,
Brielmeier, M.,
Bornkamm, G. W.,
and Behne, D.
(2001)
FASEB J.
15,
1236-1238 |
6. | Behne, D., and Kyriakopoulos, A. (2001) Annu. Rev. Nutr. 21, 453-473[CrossRef][Medline] [Order article via Infotrieve] |
7. | Roveri, A., Maiorino, M., Nisii, C., and Ursini, F. (1994) Biochim. Biophys. Acta 1208, 211-221[Medline] [Order article via Infotrieve] |
8. | Knopp, E. A., Arndt, T. L., Eng, K. L., Caldwell, M., LeBoeuf, R. C., Deeb, S. S., and O'Brien, K. D. (1999) Mamm. Genome 10, 601-605[CrossRef][Medline] [Order article via Infotrieve] |
9. | Dreher, I., Schmutzler, C., Jakob, F., and Kohrle, J. (1997) J. Trace Elem. Med. Biol. 11, 83-91[Medline] [Order article via Infotrieve] |
10. |
Ursini, F.,
Heim, S.,
Kiess, M.,
Maiorino, M.,
Roveri, A.,
Wissing, J.,
and Flohe, L.
(1999)
Science
285,
1393-1396 |
11. |
Cheng, W. H., Ho, Y. S.,
and Ross, D. A.
(1997)
J. Nutr.
127,
1445-1450 |
12. | Yamamoto, Y., Brodsky, M. H., Baker, J. C., and Ames, B. N. (1987) Anal. Biochem. 160, 7-13[Medline] [Order article via Infotrieve] |
13. |
Roveri, A.,
Casasco, A.,
Maiorino, M.,
Dalan, P.,
Calligaro, A.,
and Ursini, F.
(1992)
J. Biol. Chem.
267,
6142-6146 |
14. | Conz, P. A., Bevilacqua, P. A., La, Greca, G., Danieli, D., Rodighiero, M. P., Cavarretta, L., Maiorino, M., Roveri, A., and Ursini, F. (1993) Exp. Nephrol. 1, 376-378[Medline] [Order article via Infotrieve] |
15. | Linia, L., Majello, B., and de Luca, P. (1997) Int. J. Biochem. Cell Biol. 29, 1313-1323[CrossRef][Medline] [Order article via Infotrieve] |
16. | Adamson, E. D., and Mercola, D. (2002) Tumor Biol. 23, 23-93 |
17. |
Yomogida, K.,
Ohtani, H.,
Harigae, H.,
Ito, E.,
Nishimune, Y.,
Engel, J. D.,
and Yamamoto, M.
(1994)
Development
120,
1759-1766 |
18. | Schoonjans, K., Brendel, C., Mangelsdorf, D., and Auwerx, J. (2000) Biochim. Biophys. Acta 1529, 114-125[Medline] [Order article via Infotrieve] |
19. | Jaiswal, A. S., and Narayan, S. (2001) J. Cell. Biochem. 81, 262-277[CrossRef][Medline] [Order article via Infotrieve] |
20. | Sassone-Corsi, P. (1998) Int. J. Biochem. Cell Biol. 30, 27-38[CrossRef][Medline] [Order article via Infotrieve] |
21. | Maiorino, M., Thomas, J. P., Girotti, A. W., and Ursini, F. (1991) Free Radical Res. Commun. 12-13, 131-135 |
22. | Gille, L., Staniek, K., and Nohl, H. (2001) Free Radical Biol. Med. 30, 865-876[CrossRef][Medline] [Order article via Infotrieve] |
23. | Gabbita, S. P., Butterfield, D. A., Hensley, K., Shaw, W., and Carney, J. M. (1997) Free Radical Biol. Med. 23, 191-201[CrossRef][Medline] [Order article via Infotrieve] |
24. |
Standiford, D. M.,
Sun, W. T.,
Davis, M. B.,
and Emerson, C. P.
(2001)
Genetics
157,
259-271 |
25. |
Zabel, M. D.,
Byrne, B. L.,
Weis, J. J.,
and Weis, J. H.
(1998)
J. Immunol.
165,
4437-4445 |
26. | Zhai, G., Iskandar, M., Barilla, K., and Romaniuk, P. J. (2001) Biochemistry 40, 2032-2040[CrossRef][Medline] [Order article via Infotrieve] |
27. | Draper, D. E. (1999) J. Mol. Biol. 293, 255-270[CrossRef][Medline] [Order article via Infotrieve] |
28. |
Gee, S. L.,
Aoyagi, K.,
Lersch, R.,
Hou, V., Wu, M.,
and Conboy, J. G.
(2000)
Blood
95,
692-699 |
29. |
Liu, H. X.,
Zhang, M.,
and Krainer, A. R.
(1998)
Genes Dev.
12,
1998-2012 |
30. | Modafferi, E. F., and Black, D. L. (1997) Mol. Cell. Biol. 17, 6537-6545[Abstract] |
31. | Ashiya, M., and Grabowski, P. J. (1997) RNA (New York) 3, 996-1015 |
32. | Tsukahara, T., Casciato, C., and Helfman, D. M. (1997) Nucleic Acids Res. 22, 2318-2325[Abstract] |
33. | Helfman, D. M., Ricci, W. M., and Finn, L. A. (1988) Genes Dev. 2, 1627-1638[Abstract] |
34. | Nasim, F. H., Spears, P. A., Hoffmann, H. M., Kuo, H. C., and Grabowski, P. J. (1990) Genes Dev. 4, 1172-1184[Abstract] |