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Department of Molecular and Integrative Physiology (L.L.H.,
M.A.F.D., J.k.C.) The University of Kansas Medical Center
Kansas City, Kansas 66160
The Department of Molecular
Genetics (M.S.) The University of Texas M.D. Anderson Cancer
Center Houston, Texas 77030
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Sertoli cells are regulated by a variety of signaling molecules that assist their differentiation and development. One of these, the pituitary hormone FSH, is an integral component of the regulatory network that forms the hypothalamic-pituitary-gonadal axis. This hormone provides important stimulus to Sertoli cells that induces proliferation, final maturation events, and synthesis of specific protein products (10, 11, 12). FSH recognizes and binds a cell surface receptor (FSHR) that serves as the communicative link between the pituitary FSH signal and gonadal response (11, 13, 14). The expression of this receptor is remarkably cell specific: it is located exclusively on Sertoli cells of the testis and granulosa cells of the ovary (11, 13, 15, 16). Because of its cell-specific properties and its role in FSH signaling, determining the mechanisms that regulate FSHR gene expression will provide important insight into the regulation of FSH response and mechanisms that govern cell-specific expression in granulosa and Sertoli cells.
Studies with transgenic mice and transient transfection have been used to examine the transcriptional mechanisms that regulate FSHR (17, 18, 19, 20). Experiments using transgenic mice revealed that 5,000 bp of 5'-flanking sequence could direct expression of a reporter gene to the gonads, implicating this region in the cell-specific expression of FSHR (18). Moreover, transient transfection analysis of deletion and block replacement mutants indicated that, within this 5,000-bp gene fragment, a single E box element, located approximately 30 bp upstream of the transcriptional start site, is the major control site for FSHR transcription (17, 18, 19). Together, these studies suggest that a promoter region that includes the E box would be sufficient for cell-specific expression and that this element is an important contributor to this process. However, identification of the upstream stimulatory factor (USF) proteins, USF1 and USF2, as the major Sertoli cell E box-binding complex obscured this interpretation, as these proteins are ubiquitous and therefore insufficient, by themselves, to direct cell-specific expression (17, 19). Thus, if the E box and the USF proteins are involved, other mechanisms, such as cell-specific modifications or the use of specific coactivators, need to be invoked. Alternatively, proteins other than USF1 or USF2, not observed in the binding assays, may interact with this element and help direct cell-specific expression. To address these possible mechanisms, more direct evidence for USF regulation of FSHR is needed as well as a better-defined region that directs cell-specific expression. Through cotransfection analysis of wild-type and mutant USF proteins, we provide data that showed USF1 and USF2 activated FSHR transcription and that this activation required several domains within the amino-terminal portion of the proteins. In addition, we show that occupancy of the E box in vivo was cell specific, but likely requires assistance of sequences outside of -5,000/+123 bp of the FSHR gene.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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N and U2N, efficiently bind DNA but are unable to
transactivate due to the absence of the amino-terminal transactivation
domains (24, 25). Cotransfection with increasing amounts of vectors
expressing the wild-type USF proteins showed little impact on FSHR
promoter activity (USF1 and USF2, Fig. 1A
). However, cotransfection with
increasing amounts of vector DNA expressing the amino-terminal
transactivation mutant proteins (U1N or U2N) resulted in a
dramatic reduction in FSHR promoter activity (Fig. 1B). The effects of
the N mutants were promoter specific and E box dependent, as
luciferase production directed by either the SV40 promoter (in
pGL3-Control) or a mutant FSHR promoter lacking the E box (FSHREbox)
was insensitive to cotransfection with the N mutants (Fig. 1C). In
addition, no specific effects of the USF mutants were observed on
either the thymidine kinase or Rous sarcoma virus promoters (data not
shown). The results therefore indicate that the USF N mutants
regulate the FSHR promoter in a manner that is promoter specific and
dependent on the E box.
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N mutants suggested that USF regulates FSHR expression
and that the results observed with the wild-type proteins were due to
high levels of endogenous USF proteins. The inability of the wild-type
USF proteins to regulate FSHR promoter activity did not appear to be
due to the level of protein expressed from the vectors, as both USF1
and USF2 expression was comparable to expression of the mutant proteins
as observed by Western blot analysis (Fig. 3B). However, if the E box
in the transfected promoter was already saturated with USF due to high
levels of endogenous proteins, the ability to influence promoter
activity would be compromised. To address this possibility, we used two
different FSHR promoter mutants that contained subtle changes in the E
box sequence (µ9.3 and µ9.6, Fig. 2A). As demonstrated by their diminished
ability to compete for the USF proteins in an electrophoretic mobility
shift assay (EMSA), these mutant E boxes bind the USF proteins with
lower affinity than the wild-type sequence (Fig. 2A). In addition,
basal activity of the mutant promoters was significantly less than the
wild-type (Fig. 2B and Ref. 17), revealing that the mutant promoters
are not saturated with regulatory proteins when transfected into MSC-1
cells. When these mutant constructs were cotransfected with expression
vectors for either USF1 or USF2, promoter activity increased 2 to 3
times as compared with empty vector alone (Fig. 2B). In contrast, a
mutant promoter with the entire E box mutated (µ9.2) was only
modestly influenced by cotransfection with vectors expressing the
wild-type USF proteins (Fig. 2B). Thus, under conditions where the E
box has a lower affinity for the USF proteins and the element is not
saturated with endogenous USF, USF1 and USF2 activated FSHR promoter
activity. The results in Fig. 1
further indicated that this
transcriptional activation requires a domain or domains within the
amino-terminal portion of these proteins.
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). The
wild-type USF did not inhibit promoter activity, indicating that
inhibition is not due to the sequestration of important factors away
from the promoter. Deletion of amino acids 6–40 of USF2
(U26–40) reduced promoter activity 30–40%, revealing a
transactivation domain within this region of the protein (Fig. 3A).
Removal of an additional 83 amino acids that encompasses most of exon 4
(U27–123) had no further influence on promoter activity. Deletion
of exon 4 alone also had little impact on promoter function (Fig. 3A, U2E4), while progressively larger deletions through exon 5
(constructs U27–148 through U21–199) resulted in sequentially
greater impact on promoter activity. This suggested that multiple
regions within exon 5 act to support transcriptional activity of USF2.
Removal of the USF-specific region (USR) domain, a region of
high sequence conservation among USF proteins, further decreased
promoter activity. However, removal of either exon 5 or the USR alone
(U2E5 and U2USR) failed to inhibit promoter function, indicating
that redundant transactivation domains act within the amino terminus of
USF2. Western blot analysis of cells transfected with each of the
mutants showed that the observed results were not due to differences in
expressed protein (Fig. 3B).
An important transactivation domain within the first 130 amino acids of
USF1 was also identified (Fig. 3A, U11–130), and further deletion
through the USR resulted in a modest reduction in FSHR promoter
activity. Thus, for both USF1 and USF2, multiple domains appear to
contribute to transactivation of the FSHR gene and, at least for USF2,
there is redundancy in the ability of some of these (exon 5 and USR) to
transactivate the promoter.
The E Box of the Endogenous FSHR Gene Is Occupied in Expressing
Cells and Vacant in Nonexpressing Cells
The central role of the E box in FSHR promoter
function, as assessed by transient transfection, prompted examination
of this site on the endogenous FSHR gene using in vivo
genomic footprinting. Primary cultures of rat Sertoli cells and a rat
choriocarcinoma cell line, RCHO, that does not express FSHR were
treated in vivo with dimethyl sulfate (DMS), genomic DNA was
isolated, and the methylation pattern of guanines (and to a lesser
extent adenines) was determined after piperidine cleavage and
ligation-mediated PCR. Genomic footprints generated from the gene
in vivo were compared with footprints generated from DNA of
the same source isolated before treatment with DMS (in vitro
footprint). Comparison of the in vivo and in
vitro footprints revealed that guanines within the FSHR E box
(marked by arrows in Fig. 4, A
and B) were protected from methylation in cultured Sertoli cells
treated with DMS in vivo (T lanes, Fig. 4A) when compared
with naked DNA treated with DMS in vitro (N lanes, Fig. 4A).
Densitometric analysis revealed that, in Sertoli cells, the intensity
of the E box bands was 65% lower in the in vivo DMS-treated
DNA compared with naked DNA treated with DMS. In contrast, the
intensity of the E box bands in the RCHO cells was slightly higher
(127%) with in vivo DMS treatment than with in
vitro treatment, indicating that the E box is not protected from
methylation in RCHO cells (Fig. 4A). These studies indicate that the E
box within the endogenous FSHR gene is occupied by regulatory
proteins in FSHR-expressing cells (Sertoli) but not in cells that fail
to express the receptor (RCHO). The inability to detect a footprint
over the E box in RCHO cells was not due to the lack of USF proteins,
as EMSA and antibody supershifts revealed the presence of these
proteins in this cell line (Fig. 4C).
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; -5,000/+123Cre,
-100/+123Cre). Sixteen different lines of animals were obtained (eight
lines from each construct), and the F1 progeny were examined for Cre
recombinase expression by RT-PCR analysis of RNA isolated from 10
different adult tissues using primers that span intron D of the bGH
gene (Cre6 and PolyA5; Fig. 5A). Expression of Cre mRNA was determined
by the presence of a 334-bp amplified product as compared with a 609-bp
product derived from genomic DNA, as shown for animal 527, a progeny of
founder animal 254 (Fig. 5B).
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gives the results obtained from
animals that harbor the -5,000/+123Cre transgene. The results show
that each transgenic animal expressed Cre recombinase in the testis as
well as the brain (expression is indicated as +). While expression was
restricted to these two tissues in most of the animals, various levels
of ectopic expression were observed in some lines, most notably lines
240 and 207. Table 2 gives the results
obtained from animals containing the -100/+123Cre transgene. In two
lines, 252 and 303, no transgene expression was observed. However,
three lines, 254, 289, and 296, expressed Cre only in the testis, while
the remaining three lines showed various levels of ectopic expression.
In female mice, transgene expression was observed in the ovary of each
FSHR(-5,000)Cre transgenic line examined, except for line 232, while
only lines 256, 323, and 347 of the FSHR(-100)Cre animals exhibited
ovarian transgene expression (Tables 1 and 2). Thus, despite the rather
high level of ectopic expression, especially with the -5,000-bp
construct, the results with lines 254, 289, and 296 suggested that
sequences within the first 100 bp of the promoter are sufficient for
testis-specific expression but lacked information necessary for
ovary-specific expression.
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and 2, all animals
examined expressed Cre recombinase in the testis at 30 days of age.
However, with the exception of animal 347, none of the animals examined
expressed Cre recombinase in the testis at 10 days of age, while FSHR
expression was observed in the testis of all animals (data not
shown).
The different temporal expression patterns of FSHR and Cre strongly
suggested that Cre expression in the testis was within the germ cell
population rather than in Sertoli cells. To test this possibility, Cre
recombinase expression was examined in Sertoli cells and germ cells
isolated from the testis of lines 254 and 347. After 35 rounds of
amplification in a RT-PCR reaction, a very modest signal for expressed
Cre recombinase was observed when Sertoli cell RNA from line 347 was
used as template. However, a robust signal was detected when germ cell
RNA from the same animal was used as template (Fig. 6, SC and GC lanes, respectively). This
difference was not attributed to variation in the cDNA synthesis as no
obvious difference was observed for amplification of the control mRNA
L7 (Fig. 6, bottom). No amplified product for Cre
recombinase mRNA was observed in cell preparations from negative
littermates (Fig. 6, Neg SC and GC samples). Similar results were
obtained for transgenic line 254 (Table 2). Since germ cells, in small
amounts, often contaminate Sertoli cell preparation, the weak Cre
signal is Sertoli cells most likely represents a germ cell contribution
to this RNA pool. Thus, the high degree of ectopic Cre expression and
temporal misregulation in the testis indicate that the region from
-5,000 to +123 bp of the rat FSHR gene was insufficient to properly
restrict or direct expression to Sertoli cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Studies reported herein have yielded several important findings
regarding the mechanisms governing FSHR expression. Through
cotransfection analysis, we have shown that both USF1 and USF2 activate
FSHR transcription. Although the wild-type proteins failed to
significantly activate the FSHR promoter over the level due to
endogenous transcription factors, the effect of mutant USF proteins
strongly supported the conclusion that USF is involved in FSHR
transcriptional regulation. Mutants lacking the amino- terminal
transactivation domain (N mutants) greatly inhibited FSHR promoter
function, and this inhibition was promoter specific and required the E
box (Fig. 1), demonstrating that USF binds specifically to the FSHR
promoter in vivo. The inhibitory activity of the N
mutants was attributed to a competition between the mutants and
endogenous USF proteins for binding to the E box. Thus, if proteins
other than USF were responsible for activation through the E box,
cotransfection of the wild-type USF proteins should have inhibited as
well. This conclusion is further supported by the observation that
cotransfection with expression vectors for wild-type c-Myc, another
bHLH-Zip protein that shares the same binding site requirements as USF,
inhibited FSHR promoter activity (data not shown). In addition, more
direct evidence for USF activation of the FSHR promoter was provided by
our finding that cotransfected USF1 and USF2 increased transcription
from promoters containing E box sequences having lower affinities for
the USF proteins. This also suggested that the inability of wild-type
USF to regulate the wild-type promoter was due to saturation of the E
box with endogenous USF proteins.
The transactivation domains of these important regulatory proteins were mapped using a variety of USF1 and USF2 mutants. By evaluating the ability of the mutants to inhibit promoter activity, three main transactivation domains of USF2 and two for USF1 were identified. For USF1, the first 130 amino acids appeared to be most critical for transactivation, while modest affects were observed with a sequential deletion through exon 5 and the USR. Similar to our findings, Roy et al. (24) showed that deletion of the first 130 amino acids significantly diminished transactivation by USF1, while further deletion through the USR had little effect. Luo and Sawadogo (25) also identified the amino-terminal portion of USF1 as critical for transactivation, but their data support a more prominent role for the USR. This difference may reflect the use of distinct promoters, which has been shown to influence the mechanism of USF transactivation (24, 25).
Activation of the FSHR promoter by USF2 required domains within the first 40 amino acids of the protein, within the region encoded by exon 5, and within the USR. However, both the exon 5 region and the USR appeared redundant with other sequences in the amino-terminal domain, as deletion of either one alone did not alter FSHR promoter function. Although previous studies also identified exon 5 and the USR as containing important transcriptional activation domains, several notable differences are apparent when comparing these earlier results with those observed on the FSHR promoter (25). Thus, in contrast with our results, two additional studies failed to reveal a significant role for the first 40 amino acids of USF2 in transactivation (25, 26). Interestingly, studies with USF1 identified this domain as important but found that its activity depended on the context of the promoter examined (24). Furthermore, Luo and Sawadogo found that deletion of the USR alone abolished transcriptional activation by USF2, while in our studies, function of the USR was only revealed when other amino-terminal sequences were absent (25).
Our studies also examined a USF2 construct (U2E4) that represents a
natural splice variant that is expressed at various levels relative to
the wild-type protein in different cell types (29, 30). Although no
transcriptional effect was observed with this mutant on either the FSHR
or the adenovirus major late promoter, studies on the major
histocompatibility complex (MHC) class I gene clearly identify exon 4
as encoding a critical activation domain (26). Thus, comparison of our
studies with others already published on USF transactivation domains
show an emergence of data implicating differences in the functional
activity of these proteins. For the FSHR promoter in MSC-1 cells, the
USF proteins appear to function somewhat differently than what has been
reported for the activation of either the adenovirus major late or
MHC promoters. Interestingly, earlier studies have shown that
the transactivation properties of the USF proteins are both cell type
and promoter context dependent, suggesting that these contribute to the
differences observed (24, 25, 31, 32). Currently, information on the
domains of USF proteins is limited, and a more thorough comparison on
mechanisms of USF transactivation awaits additional studies that employ
a variety of cell types and promoters.
The promoter region of the FSHR gene was examined in Sertoli cells using in vivo genomic footprinting. Like in vitro footprinting, this approach predicts that protected regions represent sites of protein/DNA interaction and are important sites of transcriptional regulation. Importantly, though, in vivo footprinting has the added advantage of examining interactions within living cells where the chromatin conformation and concentrations of regulatory proteins have not been altered. The signal obtained from the footprinted region depends on the endogenous occupancy level of the site as well as the number of expressing cells in the preparation. Analysis of the FSHR promoter region showed that the signal associated with the two guanines within the E box was significantly lower in DNA treated in vivo with DMS compared with DNA treated in vitro. Moreover, in RCHO cells, no apparent difference was observed between in vivo and in vitro footprints with respect to the extent of methylation of this site. Examination of other nonexpressing cell lines rendered the same results as the RCHO cells (data not shown). Thus, despite the presence of the USF proteins in these nonexpressing cells, they do not bind to the E box of the endogenous FSHR gene.
To delineate the region of the FSHR gene needed for cell-specific expression, we generated transgenic mice harboring two constructs containing variable amounts of 5'-flanking sequence (5,000 bp and 100 bp). Eight different lines of animals carrying the 5,000-bp promoter were examined, but none showed selective expression in testis or the correct temporal profile for the FSHR gene. These results suggested that sites outside this region are needed to appropriately restrict expression of FSHR to the gonads and to activate the gene at the appropriate time during development. Surprisingly, upon first observation, a smaller region (-100) of gene appeared to better restrict transgene expression, as three lines showed expression only in the testis. However, in these same three lines, no transgene expression was observed in the ovary, and closer inspection of the males revealed that testis expression was due to inappropriate expression in the germ cells.
In apparent contrast to our findings, previous studies had shown that 5,000 bp of the rat FSHR promoter could direct expression of a reporter gene (ß-galactosidase) to the gonads in transgenic mice (18). However, these earlier studies did not determine the cell types that expressed ß-galactosidase in testis or ovary (histochemical and immunohistochemical analysis for ß-galactosidase was inconclusive) suggesting that, like our findings, expression in the germ cells may have contributed to the majority of the signal in the gonads. In addition, the fewer number of transgenic lines (2 vs. 16) and tissues examined (6 vs. 11) and variation in the assay sensitivity (Northern blot analysis vs. RT-PCR) may likely have contributed to differences in the amount of ectopic expression observed in the two studies. Thus, although these earlier studies implicated the region from -5,000/+69 in cell-specific expression, the studies presented here suggest that earlier conclusions may be incorrect and that sequences outside this region are needed for cell- specific expression. Nonetheless, it is important to note that differences in the transgenic constructs (-5,000/+69 vs. -5,000/+123 of promoter and different reporters) may also have contributed to the differences in expression profiles.
Both FSHR and LH receptor (LHR) are members of the glycoprotein hormone receptor subfamily within the superfamily of G protein-coupled receptors (33, 34). Comparison of the gene structures for these proteins suggested that they evolved from a common ancestral gene and revealed that the promoters of each are TATA-less (28, 35, 36). LHR, like FSHR, is expressed predominantly in cells of the ovary and testis, where it is found in testicular Leydig cells and ovarian theca and granulosa cells (13). Interestingly, recent studies with transgenic mice carrying 2 kb of the murine LHR promoter revealed some remarkable similarities with our FSHR transgenic studies (37). Thus, similar to the FSHR transgenes, three of five LHR transgenic lines exhibited expression within the testis but failed to express in the ovary, while all lines showed ectopic expression in the brain. Also of notable similarity were the observations that testis expression of the LHR transgene was not seen in earlier stages of postnatal development (before 5 weeks) and that it later appeared in the germ cell population (elongating spermatids and spermatogonia). However, in contrast to our studies, the investigators did observe transgene expression within the appropriate cells (Leydig cells) of the testis, indicating that the LHR promoter could recapitulate some of the regulatory features of the endogenous gene. Thus, while both the LHR and FSHR promoters failed to properly restrict expression of the transgenes, only the LHR promoter was able to direct expression to one of the correct gonadal cells (Leydig).
The studies in transgenic mice revealed several important findings.
First, the extensive amount of ectopic expression (predominantly brain
and germ cell) as well as two silent transgenes (nos. 252 and 303)
indicated that the FSHR transgenes were strongly influenced by adjacent
DNA sequences at the position of transgene integration. In addition,
since little or no Sertoli cell expression was detected, the elements
needed to confer and/or enhance cell-specific expression were absent
from the transgene construct. Thus, additional regulatory sequences are
needed to overcome chromatin influences at the site of integration and
to enhance transcription in Sertoli cells. The transgenic mouse
studies, together with the cotransfection and in vivo
genomic footprinting data, suggest a mechanism whereby sequences
outside the region from -5,000 to +123 alter the transcriptional
capabilities of the FSHR gene and permit or assist the occupancy of the
E box by the USF proteins (Fig. 7). Thus,
in FSHR-expressing cells, proteins bound to distal regions of the gene
will likely be required for transactivation, while in nonexpressing
cells these proteins are either absent or nonfunctional on the FSHR
gene. The mechanisms involved in these transcriptional changes remain
to be determined, but may involve alterations in chromatin structure
that permit the USF proteins to bind the E box. Alternatively, proteins
bound to distal regulatory elements may directly interact with the USF
proteins and assist their interactions with the E box in the proximal
promoter. Identification of these regulatory sequences and their
binding proteins will be critical to our understanding of the
mechanisms regulating FSHR.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Transgenic Mouse Production
The 7,333- and 2,433-bp inserts of FSHR(-5,000)Cre and
FSHR(-100)Cre were excised from plasmid sequence by digestion with
SstI and XhoI and resolved by agarose gel
electrophoresis, and the fragments were isolated using Prep-A-Gene
matrix according to the manufacturers instructions (Bio-Rad Laboratories, Inc. Hercules, CA). DNA was eluted in 10
mM Tris, pH 7.5, 0.25 mM
EDTA. Transgenic founders were produced by pronuclear injection into
B6/SJL F1 zygotes performed by the Center for Reproductive Sciences
transgenic core at the University of Kansas Medical Center. The Cre
founders were back-crossed into a CD1 background for production of
transgenic offspring. All animal studies were conducted in accordance
with the principles and procedures outlined in "Guideline for Care
and Use of Experimental Animals."
Genotyping of Transgenic Mice
The genotypes of all offspring were analyzed by PCR of mouse
tail DNAs. Mouse tail cuts (
3 mm) were incubated in 20 µl lysis
buffer (50 mM Tris-HCl, pH 8.0, 20 mM NaCl, 1
mM EDTA, 1% SDS) with 40 µg proteinase-K at 55 C for
30–40 min. The mixture was briefly vortexed after 15 min. After
incubation, 178 µl of distilled water were added, and the mixture was
heated in a boiling water bath for 5 min. The sample was cooled to room
temperature, and 1.5 µl of the tail lysate were amplified by PCR (28
cycles for 40 sec at 94 C; 40 sec at 56 C; and 40 sec at 72 C) using
Cre-specific primers Cre1 5'-CTGGTCGAAATCAGTGCGTTC-3' and Cre2
5'-TTACCGGTCGATGCAACGAGT-3'. Positive animals were identified by the
presence of a 393-bp amplified product as observed by agarose gel
electrophoresis.
Analysis of Transgene Expression
F1 transgenic mice positive for either FSHR(-5,000)Cre or
FSHR(-100)Cre transgenes were killed and total RNA was isolated from
various tissues using TRIZOL reagent according to manufacturer’s
procedures (Life Technologies, Inc., Gaithersburg, MD).
Tissue-specific expression of the transgenes was evaluated using
RT-PCR. Complementary DNA was generated using Superscript reverse
transcriptase (Life Technologies, Inc.), 2 µg total RNA,
and 0.25 µg oligo-dT. Complementary DNA synthesis was performed in
both the presence and absence of reverse transcriptase. The cDNA was
used as a template in a PCR with intron-spanning primers specific for
the Cre recombinase and exon 5 of the bGH gene (upstream primer: Cre6,
5'-CTGGATAGTGAAACAG-GG-3'; and downstream primer: polyA5,
5'-GTACGTCTC-CGT-CTTAT-3'). This primer set was
used to distinguish between genomic DNA (a contaminant in the RNA
preparation) and the expressed transgene. In addition, each cDNA was
examined for the presence of the ribosomal protein L7 to confirm cDNA
synthesis (upstream primer: L7.1,
5'-GGAAAG-GCAAGGAGGAAGCA-3'; downstream primer: L7.2,
TCCTCCATGCAGATGATGC). Amplified products were examined by agarose gel
electrophoresis in which amplification of unprocessed transgene from
genomic DNA resulted in the presence of a 609-bp DNA fragment, while
cDNA generated from processed mRNA resulted in the amplification of a
334-bp fragment. In studies examining Cre recombinase expression in
mouse Sertoli cells and germ cells, samples were also examined for the
expression of FSHR using mouse-specific primers to the FSHR (mFSHR2
5'-GGGGAAGCTTTTGGAGGTAATAGAGGCAGAT-3' and mFSHR3
5'-GGGGTCTAGAGCCTTAAAATAGACTTGTTGCA-3'). In these samples, FSHR
was detected only in the Sertoli cell preparations.
Isolation of Mouse Sertoli and Germ Cells
Seminiferous tubules were prepared from mouse testes (from
either 27- or 50-day- old animals) as described elsewhere but with
slight modification (38). For isolation of mouse seminiferous tubules,
the order of collagenase and trypsin treatments was reversed. Tubules
were cultured in Ham’s F12 media containing 5% FBS, 1.47
mM L-glutamine, 1.5 mM HEPES, 1%
penicillin/streptomycin, and 3 µg/ml cytosine arabinoside at 37 C in
5% CO2. After 5 days, germ cells were collected
by dislodging the loosely adherent germ cells into the media with
several rounds of pipeting. The collected media were then subjected to
centrifugation (200 x g for 6 min) to pellet the germ
cells. Sertoli cells were collected 2–4 days later.
Transfection and Enzyme Analysis
The mouse Sertoli cell line MSC-1 (39) was seeded onto six-well
plates (35-mm/well) at a density of 250,000 cells per well. Unless
otherwise stated, cells were transfected with 1 µg luciferase
reporter, 50 ng pRL-TK, and 0.5 µg expression vector using 5 µl
lipofectamine reagent (Life Technologies, Inc.), as
described previously (17). pRL-TK expresses Renilla
luciferase from the herpes simplex virus thymidine kinase promoter and
was included to control for transfection efficiency (Promega Corp. Madison, WI). Sixty hours after transfection, the cells
were lysed and assayed for both firefly and Renilla
luciferase activities using the Dual-Luciferase Reporter Assay System
(Promega Corp.). Specifics of the transfection procedure
are described elsewhere (17, 40). Data were averaged over a minimum of
three independent experiments.
In Vivo Footprinting
Primary rat Sertoli cells were cultured on 150-mm culture dishes
as described previously (38). RCHO cells were grown as described (41).
Cells were treated with a concentration of 0.1% DMS
(Sigma-Aldrich Corp., St. Louis, MO) in prewarmed media
for 2 min, washed three times with PBS, and lysed to isolate genomic
DNA. Genomic DNA was isolated and extracted as described elsewhere
(42). Both in vivo DMS-treated DNA and control in
vitro treated DNA were prepared simultaneously and subsequently
cleaved at methylated guanines using piperidine (Sigma-Aldrich Corp.) diluted 1:10 in water. Piperidine was removed by repeated
DNA precipitation and resuspension in H2O. DNA
samples were resuspended in water and the concentration determined by
reading the absorbency at 260 nm. Isolated genomic DNA (2 µg) was
analyzed by ligand-mediated (LM)-PCR as described elsewhere
(42). Oligodeoxynucleotides used for the ligated linker are published
elsewhere (42). LM-PCR primers to the FSHR gene, designed according to
parameters and specification described in Ref. 42 are as follows:
+168 bp to +143 bp, Tm=60.7 C,
(AS) 5'-d (GACACAGCCAGTGATGACATCCAGAT)-3'; +151 bp to +123 bp,
Tm = 64.3 C, (AS)
5'-d(CATCCAGATCCCGTGCCCAAGAATGC)-3'; +151 bp to +118 bp,
Tm = 70.1 C, (AS)
5'-d(CATCCAGATCCCGTGCCCAAGAATGCCAGCAAGG)-3'. The first primer
extends a FSHR-specific promoter fragment. The second primer, in
combination with the linker primer (described in Ref. 42), amplifies
the fragments. The last primer is labeled to a high specific activity
with P32-ATP using T4 kinase (New England Biolabs, Inc., Beverly, MA) and used to label amplified
fragments. Amplified products were fractionated on a 8% denaturing
gel, dried, and analyzed by autoradiography. Optical densities were
quantified for autoradiography bands using Gel-Pro Analyzer image
analysis software (Media Cybernetics, Silver Spring, MD). To adjust for
loading differences across the gel, optical densities of seven
different bands were normalized, and the relative intensity of the E
box was calculated for each. The average of the seven values was then
used to determine the change in the optical density of the E box bands
between treated and naked DNA samples.
Preparation of Nuclear Extracts and EMSA
To prepare nuclear extracts, cells were washed with ice-cold
buffer (25 mM HEPES, pH 7.4, 1 mM
dithiothreitol (DTT), 1.5 mM EDTA, 10% glycerol) and
scraped from plates into the above buffer with 0.5 mM
phenylmethylsulfonyl fluoride (PMSF) added. Cells were lysed using 30
strokes of a Dounce homogenizer (B pestle), and the nuclei were
pelleted by centrifugation (16,000 x g) for 1 min. The
supernatant was removed and nuclei resuspended in extraction buffer (25
mM HEPES, pH 7.9, 1 mM DTT,
1.5 mM EDTA, 10% glycerol, and 0.5
M KCl). Nuclei were extracted on ice for 10
min, and then frozen on dry ice. Nuclei were then thawed and
centrifuged (85,000 x g) for 6 min. Supernatants were
immediately aliquoted and placed at -80 C. Protein concentration was
determined by the BCA method (Pierce Chemical Co.,
Rockford, IL) using BSA as a standard.
For EMSAs, nuclear extracts (6–10 µg protein) were incubated with 25 fmol of radiolabeled double-stranded oligonucleotide in the presence of 10 mM HEPES, pH 7.9, 3 mM MgCl2, 30 mM KCl, 0.5 mM DTT, 12% glycerol, 0.6 mM EDTA, 0.2 mM PMSF, 50 ng salmon sperm DNA, 1 µg dIdC, 10 µM ZnCl, and 1 µg/ml BSA in a 20 µl reaction volume as described elsewhere (17). Addition of competitors or antibodies to the reaction immediately preceded the addition of extract. Reactions were incubated on ice for 10 min before addition of probe, and then an additional 30 min on ice before being loaded onto the gel unless otherwise noted. Protein-DNA complexes were resolved on a 4% nondenaturing polyacrylamide gel (acrylamide:bis-acrylamide = 40:1) run in 25 mM Tris (pH 8.5), 190 mM glycine at 250 V for 1.5 h at 4 C. Gels were dried and analyzed by autoradiography. Antibodies for USF1 (C-20)X, USF2 (C-20)X, and c-Myc (N-262)X were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). These antibodies are supplied as rabbit polyclonal IgG and were used directly as supplied by the manufacturer at 1 µg IgG per binding reaction.
Western Blot Analysis
MSC-1 cells were transfected with 1.5 58 g of each USF
mutant as described above. Whole cell extracts were resolved on 10%
SDS-polyacrylamide gels (acrylamide:bis-acrylamide = 30:0.8) with
the discontinuous buffer formulation of Laemmli (43) and transferred to
nitrocellulose membranes (Bio-Rad Laboratories, Inc.
Hercules, CA) using a Mini Trans-Blot Electrophoretic Transfer Cell
(Bio-Rad Laboratories, Inc. Hercules, CA). Membrane-bound
proteins were probed overnight at 4 C with anti-USF1 or anti-USF2
antibodies, diluted 1:10,000 in TBST (15 mM Tris-HCl, pH
7.4, 150 mM NaCl, 0.05% Tween-20), and then subsequently
incubated with goat antirabbit horseradish peroxidase- conjugated
antibody for 90 min at room temperature. Specific protein complexes
were visualized with the enhanced chemiluminescence (ECL) system
(Amersham Pharmacia Biotech, Arlington Heights, IL).
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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This work was supported in part by NIH Grants R29HD-3521701A1 and R03HD-35871 (to L.L.H.), RO1CA-79578 (to M. S.), and National Research Service Award F32 HD-08500 (to M.F.D.).
Received for publication March 13, 2000. Revision received June 23, 2000. Accepted for publication August 9, 2000.
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