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INTRODUCTION |
The loss of a normal cellular protein,
FMRP,1 causes fragile X
syndrome, one of the most common forms of mental retardation (MR). FMRP
is a RNA-binding protein that contains two hnRNP K-homology (KH)
binding domains and an arginine-glycine-rich region that resembles an
RGG box (1, 2). Several studies indicate that both the KH2
domain and the arginine-glycine-rich region likely play a role in RNA
binding (1, 3-6), the latter interaction being mediated by a G quartet
(7). FMRP associates with polyribosomes via a mRNP particle (8, 9), and
it has been proposed to regulate gene expression post-transcriptionally
(5, 10-14). Mammalian FMRPs inhibit mRNA translation in
vitro at nanomolar concentrations in both rabbit reticulocyte
lysates (15) and in microinjected Xenopus oocytes (16).
These data suggest that translational repression may be an in
vivo function of FMRP. Indeed, the Drosophila homolog
of FMRP, dFMR1, was found to bind and negatively regulate
futsch mRNA (17).
Recent studies have begun to delineate the mRNAs that mammalian
FMRPs interact with in vivo. These studies have taken one of
two forms. On the one hand, potential FMRP target mRNAs have been
identified solely on the basis of their ability to bind to purified
recombinant FMRP (15, 16) or cell-free produced FMRP (1, 3).
Notwithstanding, it has not been determined whether any of these
mRNAs bind to FMRP in vivo. On the other hand,
mRNAs, including FMR1 mRNA, which associate with
FMRP-containing mRNPs have also been isolated from cultured cells (10,
18). However, although these messages require FMRP in the mRNP for
their association, it has not been demonstrated that they bind solely
to it. Using the former methodology, we isolated a subset of mRNAs
derived from normal adult brain that bind human FMRP (hFMRP) in
vitro (3). During the course of this investigation we also tested a number of other mRNAs for their ability to interact with hFMRP. One of these mRNAs was Xenopus elongation factor 1A
(xEF-1A). In the present paper we demonstrate that xEF-1A mRNA
binds to recombinant and cell-free produced hFMRP in vitro.
Furthermore, we show that hFMRP inhibits EF-1A mRNA translation in
cultured PC12 and COS-7 cells and that the loss of hFMRP in fragile X
lymphoblastoid cells derepresses human EF-1A (hEF-1A) mRNA translation.
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EXPERIMENTAL PROCEDURES |
Antibodies--
FMRP mAb 2160 and normal mouse serum were
purchased from Chemicon. FXR1 (Y-19) mAb and FXR2 (S-16) mAb were
obtained from Santa Cruz. Dynamin mAb and EF-1A mAb were purchased from
Upstate Biotechnology. Hsp70cP mAb (HSP-820) was obtained from
StressGen. Horseradish peroxidase-conjugated secondary antibodies were
purchased from KPL and Santa Cruz.
Buffers--
RNA binding buffer 1 contained 50 mM
Tris-HCl, pH 7.0, 2 mM MgCl2, and 150 mM NaCl. Buffer 2 contained 20 mM Hepes, pH
7.9, 2 mM MgCl2, 70 mM
NH4Cl, 0.2% IGEPAL CA630, and 50 mg/ml yeast tRNA. FMRP
purification buffer, buffer 3, contained 10 mM Hepes, pH
7.9, 300 mM NaCl, 1% Triton X-100, 1 mM
phenylmethylsulfonyl fluoride, 5 mM
-mercaptoethanol,
and 20 mM imidazole. TAE buffer contained 40 mM
Tris acetate and 1 mM EDTA. TMK buffer contained 50 mM Tris-HCl, pH 7.6, 10 mM MgCl2,
and 25 mM KCl.
Plasmid Clones--
pET21A-hFMRP and pET21A-I304N, encoding
hFMRP and the corresponding I304N point mutant, were gifts from
Dr. Bernhard Laggerbauer, Max Planck Institute for Biochemistry,
Germany. Each vector generates full-length FMRP with an N-terminal
His tag. pTRI-XEF, encoding Xenopus EF-1A, was obtained from
Ambion Laboratories. pAPP-695 and pT7-Control, encoding
APP695 and a 1.4-kb
-HindIII/EcoRI RNA, respectively, have been
described previously (3). pSF2-hFMRP, encoding human FMRP,
was a gift from Dr. Ben Oostra, Erasmus University, Rotterdam, The
Netherlands. pHA-hFXR1P, encoding human FXR1P, was a gift from Dr.
Gideon Dreyfuss, University of Pennsylvania.
hFMRP Production for RNA Binding Studies--
Full-length and
truncated 35S-hFMRPs were prepared by coupled in
vitro transcription translation (3).
Recombinant hFMRPs were expressed in Escherichia coli BL21
from pET21A-FMRP and pET21A-I304N (16, 19). Briefly, transformed E. coli BL21 were grown at 37 °C to 1.0 A560 in LB-Amp100 medium. 1 mM isopropyl-1-thio-
-D-galactopyranoside was
added, and the cells were grown at 30 °C overnight. Proteins were
extracted from cell pellets using B-PerTM supplemented with 300 mM NaCl, 20 mM imidazole, 5 mM
-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride, and 1 × CompleteTM protease inhibitors and purified on
nickel-nitrilotriacetic acid resin, preequilibrated with buffer 3. Bound protein was eluted with buffer 3 plus 230 mM
imidazole. hFMRP production (68-70 kDa) was confirmed by Western
blotting, and its purity was determined by Coomassie Blue staining. The
micro-BCA assay was used to determine protein concentration (20).
In Vitro RNA Target Production--
RNAs were produced from
linearized plasmids by in vitro transcription. The
transcripts were purified on QuickSpinTM columns and quantified by
UV-visible spectrophotometry at 260 and 280 nm (3).
Agarose Gel Electrophoretic Shift Assay (AGESA)--
Purified
recombinant hFMRP was bound to RNA at room temperature. Briefly, the
recombinant protein was preincubated for 10 min in buffer 1 supplemented with 0.25 mg/ml tRNA and 0.25 mg/ml ultrapure bovine serum
albumin. Subsequently, RNA was added and incubated for an additional 20 min. hFMRP·RNA complexes were then resolved on 1% TAE agarose
gels (50-60 V, 50 mA for 1-2 h) and visualized by ethidium bromide
staining. Kd values were determined from
titrations of recombinant proteins with fixed amounts of in
vitro transcribed target RNAs. The percent complex formation was
measured from scanned images of the AGESAs using IPLab Gel software
(Signal Analytics Corp.) and plotted versus the amount of
recombinant protein input into the reaction. A molecular mass of
69,000 Da was used to calculate the molar amount of hFMRP (12). The
data were fit using a nonlinear curve-fitting program Kaleidograph software (Synergy Software) (21).
To verify specific complex formation, bands from different regions of a
gel shift experiment were excised and boiled for 5 min in SDS buffer
(22). The resulting extracts were blotted and probed with FMRP mAb 2160.
Affinity Capture and Homoribopolymer Binding
Assays--
35S-Labeled hFMRP, hFMRP
RGG,
hFMRP
KH2, and hFMRP
RNB binding to
biotinylated xEF-1A RNA was measured by affinity capture using
SoftLinkTM resin in buffer 2 (3). Nonspecific binding, determined by
comparison with reactions without in vitro transcribed RNA
or nonbiotinylated xEF-1A RNA, was negligible. RNA binding was
quantified using IPLab Gel software. The percent binding was calculated
as shown in Equation 1.
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(Eq. 1)
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This corrects for load differences between the bound and unbound
fractions. For the studies presented in Fig. 2, B and
C, in which both full-length and incomplete proteins were
produced, only the full-length protein was quantified. The -fold
decrease in biotinylated xEF-1A RNA binding between full-length
wild-type and full-length truncated hFMRPs was calculated as
shown in Equation 2.
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(Eq. 2)
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xEF-1 RNA competition with homoribopolymer resins (poly(rG), and
poly(rU)) was carried out with a 10-fold molar excess of xEF-1A RNA
over poly(rN), a 2-fold molar excess of soluble poly(rG), or a 2-fold
molar excess of soluble poly(rU). Briefly, 35S-hFMRP was
incubated in buffer 2 containing 3 µg of poly(rN) resin and either
xEF-1A RNA or soluble poly(rN) (1 h, 4 °C). Unbound 35S-hFMRP was removed; the resins were then washed with 40 column volumes of buffer 2. Bound 35S-hFMRP was eluted by
boiling in SDS buffer. Biotinylated xEF-1A RNA competition with
nonbiotinylated target mRNAs was carried out using the SoftLinkTM
affinity capture protocol described above. A 5-fold molar excess of
nonbiotinylated target mRNA over biotinylated xEF-1A mRNA was
added to the reaction before adding 35S-hFMRP. Competition
was expressed as the ratio of the percentage of biotinylated xEF-1A
mRNA bound to hFMRP in the presence of competitor mRNA to the
percentage of biotinylated xEF-1A mRNA bound to hFMRP in the
absence of competitor mRNA.
Cell Culture--
Undifferentiated PC12 cells were grown at
37 °C in 5% CO2 and maintained in Dulbecco's modified
Eagle's medium supplemented with 10% fetal bovine serum. COS-7 cells
were cultured in RPMI supplemented with 5% fetal calf serum and 2 mM glutamine. Cells (3 × 105/35-mm dish)
were transfected with 1 µg of pSF2-hFMRP or with 1 µg
of control DNAs (pET21A-hFMRP, pHA-hFXR1P). 24, 48, 72, or 96 h
later the cells were harvested and used to prepare total RNA or total
proteins (3, 23). The transfection efficiency for each experiment
varied between 20 and 60%, but within a particular experiment the
efficiency was uniform. Fragile X and normal lymphoblastoid cell lines
were cultured in RPMI supplemented with 10% fetal bovine serum.
Gene Expression in Cultured Cells--
Northern blotting was
carried out as described previously (23). Probes were prepared by
amplifying the first 871 bases of human FMR1 cDNA from
pSF2-hFMRP DNA and the entire xEF-1A coding sequence from
pTRI-XEF DNA (86.7% identity to rat EF-1A mRNA). cDNAs were
random prime labeled with [
-32P]dCTP and desalted on
G-50 QuickSpinTM columns before hybridization.
Western blotting was performed as described previously (24). FMRP mAb
2160 was used at a 1:10,000 dilution; under these conditions the
antibody preferentially detects hFMRP, therefore it is not possible to
ascertain whether transient transfection results in FMRP
overexpression. HSP-820, EF-1A, and dynamin mAbs were used at a 1:5,000
dilution. Blots were blocked for 1 h at room temperature in
phosphate-buffered saline supplemented with 3% non-fat dry milk and
probed overnight in fresh buffer with the corresponding primary
antibody at 4 °C. Blots were developed using LumiGlo. Horseradish
peroxidase-conjugated goat anti-mouse secondary antibody was used at a
1:5,000 dilution. FXR1 (Y-19) antibody and FXR2 (S-16) antibodies were
used at a 1:100 dilution. Blots were blocked and probed as above and
then developed using the manufacturer's washing procedure
(www.scbt.com; Research Applications). Horseradish
peroxidase-conjugated bovine anti-goat secondary antibody was
used at a 1:2,000 dilution. Blots were probed simultaneously with two
different antibodies, an internal control antibody such as dynamin or
HSP-820 and an antibody directed to the protein of interest. Blots were
quantified from scanned images; the ratio of protein to Hsp70cP was
used to normalize all data.
Immunoprecipitation Analysis--
PC12 cells were transfected
with pSF2-hFMRP or pET21A-hFMRP. 48 h
post-transfection the cells were scraped in 1.0 ml of diethyl pyrocarbonate-treated 1 × phosphate-buffered saline and pelleted by centrifugation. The pellet was washed twice with ice-cold diethyl pyrocarbonate-treated 1 × phosphate-buffered saline. All
subsequent steps were carried out at 4 °C. Pellets were lysed with
100 µl of buffer 1 supplemented with 1% IGEPAL CA630. The lysates
(50 µl) were precleared for 3.5 h with 30 µl of protein A/G
that was pretreated with 30 µl of normal mouse serum, 10 µl of
RNAsin, 20 µl of 50 × Complete protease inhibitors, and 500 µl of buffer 1. The precleared lysates were immunoprecipitated
overnight with 30 µl of FMRP mAb 2160-coupled protein A/G beads.
After a 10-min 3,000 × g spin, proteins or RNA was
extracted from the supernatants and pellets. Supernatant proteins were
prepared by adding 3 × SDS sample buffer to the supernatant
(250:750 µl ratio). Immunoprecipitated proteins were prepared by
adding 300 µl of 1 × SDS sample buffer to each pellet. Total
RNA was extracted from the immunoprecipitant using 1 ml of
TRI-ReagentTM. The final RNA pellet was dissolved in 25 µl of
diethyl pyrocarbonate-treated H2O and used to
prepared first strand cDNA. cDNAs were amplified using
rat-specific EF-1A-specific primers (5'-ATATTATCCCTAACACCTGCC,
5'-GGTCTCAAAATTCTGTGACAG) that amplify a 259-bp fragment from bases
1464 to 1723 of rat EF-1A mRNA, accession no. X61043. FMR1-specific
primers (set A: 5'-GGCTAGAAGCTTTCTGGA, 5'-GTGAATGGAGTACCCTAA) were used
to amplify a 1,023-bp fragment from bases 945 to 1968 of mouse fmr1
mRNA, accession no. L23971; whereas set B (5'-GGCTAGAAGCTTTCTGGA,
5'-ACGTGGAGGAGGCTTCAAAGGAAA) amplifies a hFMR1-specific 833-bp
fragment from bases 831 to 1,644 of human FMR1 mRNA, accession no.
NM_002024.
Polyribosome Analysis--
Polyribosomes were prepared by
pelleting PC12 lysates through 50% sucrose pads (25). Briefly, PC12
cells (5 × 107) were transfected with
pSF2-hFMRP or pET21A-hFMRP. 24 h later the cells were
scraped from the dishes in ice-cold TMK buffer and pelleted. The
pellets were lysed in 200 µl of TMK buffer supplemented with 1%
IGEPAL CA630. 50-µl aliquots were loaded on 50% sucrose pads
containing 500 µl of TMK buffer and centrifuged at 50,000 rpm for
20 h. The resulting polyribosome pellets were washed with 200 µl
of TMK buffer and then extracted with 50 µl of 1 × SDS sample
buffer. In some cases, the lysates were treated for 5 min at 37 °C
with 25 mM EDTA to disrupt the polyribosomes (8,
25-27).
RNA Motif Analysis--
A minimal sequence WGGN1-4
WGGN1-4 WGGN1-4 WGGN0-6 was used
to assess EF-1A mRNA and individual FMRP-target mRNAs (1, 3)
for the potential presence of G quartet structures. Candidates
containing such sequences were then folded into secondary structure
models using Mfold (//bioinfo.math.rpi.edu/) or RNABOB to determine
whether a requisite hairpin stem-loop S6
NWGGN1-4 WGGN1-4 WGGN1-4
WGGN0-6NS6 formed (6).
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RESULTS |
FMRP Binds EF-1A in Vitro--
Several studies using FMR1 mRNA
have been undertaken to define the RNA motif that FMRP recognizes (3,
7, 28). The results suggest that FMRP may bind to multiple
regions of a mRNA (19); thus, in assessing FMRP target mRNA
binding, an assay that measures interactions of large mRNAs with
FMRP is necessary. To do this, we modified a nondenaturing AGESA (30)
so that unlabeled in vitro transcribed mRNAs and
mRNA·protein complexes could be visualized by ethidium bromide
staining. Fig. 1A illustrates
results obtained by incubating the 1.9-kb 3'-untranslated region of
human FMR1 mRNA (FMR1 3'-UTR) or a 1.4-kb lambda RNA fragment
(
-control RNA) in the presence or absence of purified recombinant
human FMRP (hFMRP). As shown in lane 1, FMR1 3'-UTR mRNA
migrated as two bands in the absence of hFMRP, indicating that the RNA
resides in two conformational states (31). These two conformers
coalesced into a uniquely migrating single band in the presence of
hFMRP (lane 2). Although the shift was small, the bands were
completely resolved. In contrast, the shift did not occur when the
recombinant protein was added to
-control RNA (lanes 3 and 4). This was expected because FMR1 3'-UTR mRNA binds
specifically to FMRP (1, 3, 7, 28), whereas the
-control RNA does
not (1, 3).

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Fig. 1.
Resolution of hFMRP·RNA complexes by
AGESA. A, specific hFMRP·RNA complexes form in the
presence of 50 ng of purified recombinant hFMRP and 0.1 µg of hFMR1
3'-UTR RNA (lane 2) which are resolved from unbound hFMR1
3'-UTR RNA (lane 1). These complexes (marked by an
asterisk) form in the presence of excess tRNA; a nonbinding
RNA of similar length (3) does not form complexes under
identical conditions (lanes 3 and 4).
B, 0.1 µg of xEF-1A mRNA binds to 50 ng of hFMRP
in vitro. Specific hFMRP·xEF-1A mRNA complexes
(lane 3) and hFMRP-I304N·xEF-1A complexes (lane
4) are observed under the same conditions as A. The
complexes are fully resolved from xEF-1A mRNA alone (lane
1) or xEF-1A with 2.5 µg of bovine serum albumin and 2.5 µg of
tRNA (lane 2). Complex formation is disrupted by increasing
the ionic strength of the binding buffer to 0.5 M
(lanes 5 and 6) or by prior denaturation of hFMRP
or hFMRP-I304N (lanes 7 and 8). Recombinant
proteins do not contain residual RNA (lanes 9 and
10). The effect of added salt (lanes 13-15) on
binding is shown in lanes 3 and 12. C,
binding 0-50 ng of hFMRP to 0.1 µg of xEF-1A mRNA is saturable,
and quantitative differences are observed between wild-type and I304N
mutant hFMRP. The mean values for two independent experiments are
plotted. D, 10 ng of hFMRP is specifically isolated from 0.5 µg of the xEF-1A mRNA-shifted bands. Bands 1-5 (left
panel) were excised and proteins extracted. The extracts 1-5
(right panel) were probed with FMRP mAb 2160 on Western
blots.
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Although these data are consistent with the formation of an
hFMRP·FMR1 3'-UTR complex, several control experiments were performed to confirm this observation. To our surprise, another RNA, EF-1A from
Xenopus (xEF-1A mRNA, 1.6 kb), displayed the same
feature as FMR1 3'-UTR mRNA in AGESA. As shown in Fig.
1B, the two xEF-1A mRNA conformers (lane 1)
coalesced into a uniquely migrating single band in the presence of
hFMRP (lane 3). This result was recapitulated when we used
hFMRP-I304N, containing the KH2 RNA binding domain mutation
I304N (lane 4). This mutant was of particular interest because it is associated with exceptionally severe mental retardation (32). Because NaCl concentrations above 0.25 M decrease the FMRP affinity for RNA we also examined the effect 0.5 M
NaCl had on xEF-1A mRNA binding to hFMRP and hFMRP-I304N. Fig.
1B, lanes 5 and 6, shows that the band
shift seen in lanes 3 and 4 was completely abrogated; lesser decreases were observed at lower salt concentrations (lanes 11-15). The RNA mobility shift was also lost when
the recombinant proteins were heat denatured at 65 °C for 5 min
before adding xEF-1A mRNA (lanes 7 and 8).
These data indicate that more than the mere presence of the recombinant
hFMRPs is required for the shift. In addition, the fact that 2.5 µg
of recombinant bovine serum albumin failed to produce a shift (compare
lanes 1 and 2) indicates that the response was
not due simply to added native protein. Finally, lanes 9 and
10 demonstrate that neither recombinant protein preparation
contained large molecular mass nucleic acid, indicating that the
shifted bands contained xEF-1A mRNA.
We then used AGESA to define further the binding properties of hFMRP
and xEF-1A mRNA. Fig. 1C shows that xEF-1A mRNA
binding was saturable with increasing concentrations of hFMRP or
hFMRP-I304N. The apparent Kd values for these
complexes were 3.0 and 6.1 nM, respectively. These data
corroborate the data in Fig. 1B, suggesting that hFMRP and
hFMRP-I304N binding to xEF-1A mRNA is specific.
To demonstrate further that the shifted band was an xEF-1A
mRNA·hFMRP complex, hFMRP was specifically and uniquely recovered from the putative complex. Here, xEF-1A mRNA was incubated alone or
with subsaturating amounts of purified recombinant hFMRP and subsequently resolved by AGESA. Fig. 1D shows that adding
hFMRP resulted in either the loss or decrease in the upper xEF-1A
conformers and a concurrent broadening of the lower conformer (compare
lanes 1 and 2). Five regions of this gel were
then excised and probed for the presence of hFMRP. The right
panel of Fig. 1D shows that only the xEF-1A
mRNA-shifted band contained hFMRP (lane 2). Thus, these
data demonstrate that recombinant hFMRP and xEF-1A mRNA associate
in vitro.
xEF-1A mRNA Binding Requires the C-terminal
Arginine-Glycine-rich Region--
FMRP has three RNA binding domains,
and there is no a priori basis for knowing whether one or
any combination of them interacts with a particular RNA. We have
previously used 35S-FMRP truncation mutants in affinity
capture assays to show that hFMR1 mRNA binding requires
determinants in its KH2 domain (1, 3). Therefore, we
employed this strategy to determine the domains required for binding
xEF-1A mRNA. Four different hFMRP forms were assessed: 1)
full-length hFMRP, 2) hFMRP
RGG in which the last 334 amino acids including the arginine-glycine-rich region are deleted, 3)
hFMRP
KH2 in which the arginine-glycine-rich region and
the KH2 domain are deleted, and 4) hFMRP
RNB
in which all three RNA binding domains are deleted (Fig.
2A). Fig. 2B shows
that xEF-1A mRNA bound 35S-hFMRP, recapitulating the
results of Fig. 1. 35S-hFMRP
RGG also bound
to xEF-1A mRNA, albeit with a 6.5-fold decrease in affinity
compared with full-length hFMRP. Removing the KH2 domain
(35S-hFMRP
KH2) reduced the binding further,
whereas 35S-hFMRP
RNB binding was not
detectable under the conditions of the assay. Therefore, the C-terminal
334 amino acids of hFMRP play a major role (either direct or indirect)
in binding xEF-1A mRNA; however, the KH2 domain also
influences the binding as well.

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Fig. 2.
Interaction of xEF-1A mRNA with hFMRP
protein domains. A, RNA binding domains of full-length
hFMRP, hFMRP RGG, hFMRP KH2, and
hFMRP RNB. KH domain numbering was based on sequence assignments of Lewis et al. (29).
B, biotinylated xEF-1A mRNA binding to
35S-hFMRP, 35S-hFMRP RGG,
35S-hFMRP KH2, or
35S-hFMRP RNB. Bound material was captured on
SoftLinkTM avidin resin. The unbound (U) and the bound
(B) fractions were assessed by autoradiography.
Arrows mark full-length hFMRP or the corresponding
truncation mutant. The asterisk (*) marks incomplete
or breakdown products formed during in vitro translation
(19). C, xEF-1A mRNA, target mRNAs hFMR1 CDS and
hFMR1 3'-UTR but not -control RNA, compete with biotinylated xEF-1A
mRNA in binding to 35S-hFMRP. The values for two
independent experiments are plotted. D, xEF-1A mRNA does
not compete with poly(rG) in binding to 35S-hFMRP; binding
was assessed as in B. The arrow marks
full-length hFMRP forms. The asterisk (*) marks the major
incomplete or breakdown product formed in the in vitro
translation reaction.
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hFMRP appears to use unique sets of residues in binding various parts
of FMR1 mRNA. To determine whether these residues were similar or
identical to those interacting with xEF-1A mRNA we performed
affinity capture competition assays in which 35S-hFMRP
binding to biotinylated xEF-1A RNA was competed with nonbiotinylated RNA cognates (xEF-1A, hFMR1 CDS, hFMR1 3'-UTR, or
-control). Fig.
2C shows the effect of a 5-fold molar excess of
nonbiotinylated RNA. Here, FMR1 CDS RNA and FMR1 3'-UTR RNA reduced the
amount of 35S-hFMRP bound to biotinylated xEF-1A mRNA
by 28.6 and 44.4%, respectively. In contrast,
-control RNA had no
effect. However, xEF-1A mRNA was the best competitor with a 66.7%
reduction compared with binding in the absence of competitor RNA. These
data suggest that the hFMRP residues that bind xEF-1A are similar but
not identical to those that bind FMR1 CDS RNA or FMR1 3'-UTR RNA.
Poly(rG) and poly(rU) RNA mimetics bind primarily to incompletely
overlapping residues in the last 300 amino acids of hFMRP. Therefore,
we performed a similar competition study with them. We found that a
2-fold molar excess of poly(rG) completely blocked hFMRP binding to
poly(rG) resin. In contrast, a 10-fold molar excess of xEF-1A mRNA
had no effect on the ability of hFMRP to bind to poly(rG) (Fig.
2D) or poly(rU) (not shown) resins. This suggests that the
determinants within the C-terminal region which bind to poly(rG) and
poly(rU) differ from xEF-1A mRNA, or xEF-1A mRNA binds much
more weakly to hFMRP than either homoribopolymer.
Recent studies have shown that a nucleic acid tertiary structure
element called a G quartet, whose formation is enhanced in the presence
of potassium cations, is present in several mRNAs that may interact
with FMRP (6, 7, 17, 18). Using the criteria of Darnell et
al. (6), we showed that xEF-1A mRNA lacked a perfect G quartet
motif (Table I). We then
experimentally confirmed that the hFMRP interaction with xEF-1A
mRNA was not enhanced in buffers in which we substituted 0.15 M K+ for 0.15 M Na+
(not shown). These data, then, are consistent with the hypothesis that
the interaction between hFMRP and xEF-1A mRNA does not depend on
the formation of a G quartet structure.
FMRP Inhibits EF-1A Expression in Vivo--
EF-1A mRNA is
translationally repressed by a factor that can be salt-washed from
mRNPs (33). Previous studies have also demonstrated that recombinant
hFMRP inhibits the expression of certain mRNAs (15, 16, 34). To
determine whether FMRP affects EF-1A mRNA translation, PC12 cells
were transfected with plasmids that produce human FMRP
(pSF2-hFMRP) or a nonexpressing control (pET21A-hFMRP).
hFMRP and endogenous rat EF-1A (rEF-1A) protein expression patterns
were then examined. First, transfected cell extracts were probed with
antibodies to FMRP, EF-1A, and control proteins dynamin and Hsp70cP
(Fig. 3A). Intense
hFMRP-specific staining was observed in
pSF2-hFMRP-transfected cell extracts, whereas comparatively
weaker endogenous rFMRP staining was found in pET21A-hFMRP-transfected
cell extracts. More importantly, rEF-1A levels were significantly lower
in pSF2-hFMRP extracts than in pET21A-hFMRP extracts. This
is not a pleiotrophic effect because dynamin and Hsp70cP levels were
nearly equivalent in both sets of transfected cells.

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Fig. 3.
Effects of transiently expressing hFMRP in
undifferentiated PC12 cells and COS-7 cells. A, PC12
cells were transfected with pET21A-hFMRP (lane 1) or
pSF2-hFMRP (lane 2) and harvested 24 h
after transfection. Total transfected cell proteins were blotted and
probed sequentially with FMRP mAb and dynamin mAb, followed by Hsp70cP
mAb and EF-1A mAb. B, Northern blots from an identical set
of pET21A-hFMRP- (lane 1) or pSF2-hFMRP-
(lane 2) transfected PC12 cells harvested 24 h
post-transfection. Blots were probed with an FMR1-specific cDNA or
an EF-1A-specific cDNA, as indicated. Total RNA is shown as a load
control. C, COS-7 cells were transfected with
pSF2-hFMRP (lane 1), pET21A-hFMRP (lane
2), or pHA-FXR1P (lane 3) and harvested 24 h
post-transfection. Total proteins were blotted as in A; one
blot was probed sequentially with FMRP mAb and dynamin mAb followed by
Hsp70cP mAb and EF-1A mAb; a duplicate blot was probed with dynamin mAb
and FXR1P mAb. The transfection efficiency, determined by
immunostaining for hFMRP and hFXR1P, was equivalent.
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To know whether the rEF-1A protein reduction in hFMRP-expressing cells
resulted from transcriptional or translational regulation, we performed
Northern blotting experiments using probes that specifically recognized
human FMR1 mRNA or that detected endogenous rat EF-1A mRNA
(rEF-1A mRNA). As shown in Fig. 3B, hFMR1 mRNA was
expressed abundantly in pSF2-hFMRP-transfected cells but
not in pET21A-hFMRP-transfected cells. However, there was no detectable
reduction in rEF-1A mRNA, indicating that the decrease in rEF-1A
protein levels in pSF2-hFMRP-transfected cells was not
caused by hFMRP-mediated rEF-1A mRNA instability. These data are
consistent with the hypothesis that hFMRP negatively regulates
endogenous rat EF-1A mRNA translation in vivo.
To explore further the inverse correlation between hFMRP and rEF-1A
protein expression we examined the effect hFXR1P, a homolog of hFMRP,
had on EF-1A expression. Specifically, COS-7 cells were transfected
with pSF2-hFMRP, pET21A-hFMRP, or pHA-FXR1P. Extracts from
the transfected cells were then probed with antibodies to FMRP, EF-1A,
dynamin, Hsp70cP, and hFXR1P (Fig. 3C). As
expected, hFMRP was readily detected in
pSF2-hFMRP-transfected cells, whereas the other two
transfected cell lines displayed weaker staining of endogenous monkey
FMRP. Similarly, FXR1P expression was detected preferentially in
pHA-FXR1P-transfected cells. As was the case in PC12 cells, no
discernible differences were observed in dynamin or Hsp70cP. In
contrast, EF-1A levels were much lower in
pSF2-hFMRP-transfected cells than in cells transfected with
either pET21A-hFMRP or pHA-hFXR1P. Again, EF-1A mRNA levels were
unchanged in the three different transfected cell lines (not shown).
These data corroborate those from PC12 cells, demonstrating that the
hFMRP effect on EF-1A expression does not depend on cell type.
If hFMRP suppresses EF-1A mRNA translation, then decreased hFMRP
expression should increase EF-1A protein expression. Therefore, we
transiently expressed hFMRP in PC12 cells and examined its effect on
rEF-1A, Hsp70cP, and
APP, a hFMRP-nonbinding mRNA (3), over an
extended period of time post-transfection. Fig.
4A shows an example of one
such experiment. Here, hFMRP reached its highest expression 24-48 h
after transfection and subsequently decreased to 31% of its peak value
96 h later. In contrast, Hsp70cP levels were not markedly
affected. Fig. 4B shows that as hFMRP levels decrease, a
corresponding increase in relative rEF-1A protein expression occurs,
whereas relative
APP levels were not significantly affected. In
contrast, in control extracts where endogenous rFMRP levels did not
vary, both relative rEF-1A levels and relative
APP levels were
unchanged.

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Fig. 4.
rEF-1A expression correlates inversely
with hFMRP expression in PC12 cells. PC12 cells were transfected
with pSF2-hFMRP or pET21A-hFMRP. 24, 48, 72, and 96 h
post-transfection the cells were harvested; extracted proteins were
blotted as in Fig. 3. A, hFMRP expression in
pSF2-hFMRP-transfected cells decreases over time.
Corresponding changes were not observed for Hsp70cP. B,
rEF-1A levels increase coordinately in
pSF2-hFMRP-transfected cells but are relatively constant in
pET21A-hFMRP-transfected cells. APP expression is unchanged
in pSF2-hFMRP-transfected cells and
pET21A-hFMRP-transfected cells. Western blots were probed
simultaneously with antibodies to Hsp70cP and FMRP, or Hsp70cP and
EF-1A, or Hsp70cP and APP; the FMRP:Hsp70cP, EF-1A:Hsp70cP, and
APP:Hsp70cP ratios for two independent experiments are plotted. The
amount of Hsp70cP over the period examined did not vary by more than
12%.
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To demonstrate that the relationship between hFMRP expression and
rEF-1A expression observed in Figs. 3 and 4 occurred by direct
interaction of hFMRP and rEF-1A mRNA, we immunoprecipitated hFMRP
from pSF2-hFMRP-transfected PC12 cells. We then isolated the mRNAs from the immunocomplex and determined whether rEF-1A mRNA was incorporated. As controls we immunoprecipitated
pET21A-hFMRP-transfected cells and processed
pSF2-hFMRP-transfected cells in the same manner without the
precipitating antibody. Fig.
5A shows that hFMRP was present in the pSF2-hFMRP-transfected cell supernatant and
immunoprecipitate; in contrast, hFMRP was only found in the supernatant
in the absence of FMRP mAb 2160. Finally, FMRP was not detected in
either the supernatant or the immunoprecipitate of
pET21A-hFMRP-transfected cells under the same conditions. As shown in
Fig. 3, A and C, FMRP mAb 2160 recognizes
endogenous rFMRP with much less sensitivity than hFMRP. Thus, these
results were not surprising.

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Fig. 5.
EF-1A mRNA specifically associates with
hFMRP in transiently transfected PC12 cells. PC12 cells were
transfected with pSF2-hFMRP or pET21A-hFMRP. 24 h
post-transfection the cells were harvested and lysed. FMRP was
immunoprecipitated (IP) from the lysates using FMRP mAb
2160. An identical pSF2-hFMRP transfection was processed
similarly except that FMRP mAb 2160 was omitted. Half of the sample was
used to extract proteins; half was used to extract mRNA.
A, Western blot (WB) analysis of the FMRP
distribution in immunoprecipitate (P) and supernatant
(S) fractions. 1×S and
4×P correspond to the protein loads.
B, FXR1P and FXR2P are found in
pSF2-hFMRP-transfected cell immunoprecipitates and
supernatants. C, EF-1A mRNA and FMR1 mRNA are
amplified from RNA associated with the pSF2-hFMRP
immunoprecipitate. Mouse FMRP plasmid DNA (10 ng) (lane 1)
and cDNA from total PC12 lysates, the pET21A-hFMRP
immunoprecipitate, and the pSF2-hFMRP immunoprecipitate
(lanes 2-4, respectively) were amplified with primers for
rat EF-1A mRNA (rEF-1A), rat and human FMR1 mRNA (r/hFMR1), or
human FMR1 mRNA (hFMR1), as indicated. D, FMRP
distribution in polyribosomes. PC12 cells were transfected with
pSF2-hFMRP or pET21A-hFMRP. 24 h post-transfection the
cells were harvested and lysed. Cell lysates were treated or not
treated with EDTA and polyribosomal pellets isolated. Pelleted proteins
were blotted and probed with FMRP mAb 2160 (top panels) or
assessed for protein by Coomassie Blue staining (bottom
panels).
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Next, we demonstrated that the pSF2-hFMRP-transfected PC12
cell hFMRP immunocomplex contained known FMRP-associated proteins (10,
35-38). Western blots of the supernatant and immunoprecipitate fractions were probed with either FXR1P- or FXR2P-specific antibodies; both proteins were associated with the pSF2-hFMRP
immunoprecipitate (Fig. 5B), indicating that the transiently
expressed hFMRP associates with mRNPs that contain proteins that
complex with endogenous rFMRP.
We then extracted mRNA associated with the FMRP immunoprecipitates
and amplified it with rat EF-1A-specific primers. Fig. 5C
shows that rEF-1A mRNA was amplified from cDNA generated from untransfected PC12 cell RNA and from the pSF2-hFMRP
immunoprecipitate, but not the pET21A-hFMRP immunoprecipitate. Because
FMR1 mRNA binds to FMRP, we also examined whether the
immunoprecipitate-associated mRNA contained FMR1 mRNA. Two sets
of primers were used; set A amplified a conserved 1,023-bp fragment
from different species of FMR1, and set B amplified a 833-bp human
FMR1-specific fragment. As expected, the set A primers amplified Fmr1
from as a plasmid containing full-length mouse FMRP as well as
cDNAs derived from PC12 cell total RNA and from the
pSF2-hFMRP immunoprecipitate, but not from the pET21A-hFMRP
immunoprecipitate. In contrast, set B primers exclusively amplified
hFMR1 from the pSF2-hFMRP immunoprecipitate. These results
demonstrate that rEF-1A mRNA as well as r/hFMR1 mRNA and hFMR1
mRNA associate with hFMRP in PC12 cells.
Finally, to rule out that the decreased rEF-1A expression in
pSF2-hFMRP-transfected cells occurred because heterologous
hFMRP sequestered rEF-1A mRNA in inactive mRNPs, we ascertained
whether recombinant hFMRP was incorporated into PC12 cell
polyribosomes. Western blots of pET21A-hFMRP-transfected cell
polyribosomal pellets contain a band corresponding to endogenous rFMRP
that was dissociated by EDTA (Fig. 5D). In
pSF2-hFMRP-transfected cells, a much stronger band
corresponding to recombinant hFMRP and endogenous rat FMRP was found in
the polyribosome pellet in the absence of EDTA. Again, EDTA treatment
released most of the associated FMRP. Thus, the data in Fig. 5,
C and D, demonstrate that rEF-1A mRNA
associates with hFMRP in a multiprotein mRNP complex and that
hFMRP-containing mRNPs associate with polyribosomes.
EF-1A Protein Levels Increase in Fragile X
Lymphocytes--
Fragile X syndrome results from the loss of hFMRP,
and it has been hypothesized that this loss should produce changes in
the expression of the mRNAs it interacts with (3, 39, 40). The data
obtained from transiently expressing hFMRP in PC12 and COS-7 cells
suggest that hFMRP negatively regulates EF-1A expression. If true,
human EF-1A (hEF-1A) expression should be greater in fragile X patients
than in normal individuals. To test this, full-mutation fragile X male
and normal male control lymphoblastoid cell lines were probed for EF-1A
and Hsp70cP expression. Fig.
6A shows that hEF-1A was
2.1-fold greater in fragile X cell lines than in control cell lines
(p > 0.003, analysis of variance). In contrast, hEF-1A mRNA levels were not significantly different between the two groups (Fig. 6B). Thus, hFMRP appears to regulate hEF-1A mRNA
translation in vivo negatively.

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Fig. 6.
A, EF-1A protein levels are elevated in
fragile X lymphoblastoid cells compared with control lymphoblastoid
cells. Total lymphoblastoid cell line proteins from six full-mutation
fragile X males with moderate to profound mental retardation and four
normal male controls were probed simultaneously for Hsp70cP and EF-1A
expression by Western blotting. The EF-1A:Hsp70cP ratio (relative
EF-1A) for two experiments is plotted. B, EF-1A mRNA
levels are not altered in fragile X lymphoblastoid cells. Total RNA
from control lymphoblastoid cell lines (C1-C3) and fragile
X lymphoblastoid cell lines (F1-F5) was blotted and probed
with 32P-labeled EF-1A cDNA. The rRNA load for each
sample is shown below.
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DISCUSSION |
Our data show that hFMRP binds xEF-1A mRNA in
vitro. Binding was detected both as a minimal mRNP composed of
recombinant hFMRP and xEF-1A mRNA and also as a larger mRNP (1)
using in vitro translated hFMRP. This serendipitous finding
led us to perform several experiments to confirm the observation and
then to generalize the interaction to an in vivo setting by
isolating EF-1A mRNA from hFMRP-containing immunoprecipitates. To
our knowledge this is the first demonstration that a mRNA that
binds to solely to FMRP in vitro has altered expression when
FMRP levels are modulated in vivo. Other reports, (1, 3, 12,
15, 16) have failed to demonstrate that specific in vitro
bound mRNAs bind FMRP in cells or that mRNAs associated with
FMRP-containing mRNPs bind directly to it (10, 17, 18, 35).
Using two in vitro binding assays and several different
binding conditions we found that hFMRP formed a specific complex with xEF-1A mRNA. From agarose gel shift assays conducted under
conditions used to demonstrate specific binding of the RNA binding
protein HuD (30), we observed small reproducible shifts in the
migration of a rather large xEF-1A mRNA transcript. These shifts
occurred only in the presence of native hFMRPs and were disrupted by
high salt concentrations. We also determined that the
Kd of xEF-1A mRNA and recombinant hFMRP was
~3 nM. This value is similar to Kd
values calculated for FMR1 mRNA (12) using an affinity capture assay, EMSA results for a 426-base RNA encoding part of the FMR1 CDS
(7), as well nitrocellulose filter binding experiments with 96-base
SELEX-derived RNAs (6). Thus, AGESA complements these other assays and
demonstrates that specific interactions occur between recombinant hFMRP
and mRNA in vitro.
Studies with truncated hFMRPs showed that the arginine-glycine-rich
C-terminal end was required for efficient xEF-1A mRNA binding
in vitro. Currently, we cannot differentiate whether this region interacts directly with xEF-1A mRNA or whether it merely stabilizes the domain that xEF-1A mRNA binds to. The results of Fig. 2B suggest that the KH2 domain plays at
least a small role in binding, and this is corroborated by the results
in Fig. 1C showing that the hFMRP-I304N
Kd is slightly weaker than hFMRP. Furthermore,
the protein determinants used to bind xEF-1A mRNA do not appear to
overlap completely those of FMR CDS RNA, FMR1 3'-UTR RNA (Fig.
2C), or poly(rG) (Fig. 2D), or poly(rU). Although studies using other FMRP truncations and point mutants are needed to
address this question fully our data are consistent with the view that
full-length mRNAs bind to multiple FMRP RNA binding domains.
Recent studies show that hFMRP binds to a G quartet structure in hFMR1
CDS with high affinity (7), and an in vitro selected RNA
(sc-1) with a G quartet binds to the arginine-glycine-rich-region of an
alternatively spliced hFMRP variant (6). EF-1A mRNA, however, does
not contain a perfect G quartet motif, and we determined experimentally
that the G quartet-stabilizing cation, K+ (41), did not
enhance xEF-1A mRNA binding to hFMRP. This suggests that there are
different RNA-binding determinants in xEF-1A mRNA than FMR1
mRNA or sc-1 RNA. In fact, of the 10 RNAs we have demonstrated hFMRP binds in vitro (1, 3), 5 do not contain a G quartet structure (Table I). These data extend the recently published microarray studies (18) and imply that this structure may not be as
involved in generating the fragile X phenotype as has been intimated
(42).
The effects of altering hFMRP levels in vivo were examined
in cultured cells that transiently express hFMRP and cultured
lymphoblastoid cells derived from fragile X patients that lack FMRP.
The former mimics, to a certain degree, FMRP expression during
embryogenesis where the level of FMRP rises and then levels off or
decreases in certain cells (43-46). In both cases EF-1A protein
levels, but not mRNA levels, change in response to hFMRP
expression. The effect is very specific because three other proteins,
APP, dynamin, and Hsp70cP, remain unaltered in the transfected
cells, and hFXR1P expression did not replicate the effect. These data
suggest that hFMRP regulates EF-1A mRNA translation via direct
interaction. To demonstrate this more convincingly, we
immunoprecipitated hFMRP from transiently transfected PC12 cells and
showed that rEF-1A mRNA was specifically found in the
immunoprecipitate. We also established that the expressed recombinant
hFMRP functioned normally in PC12 cells. Indeed, we showed that hFMRP
was present in polyribosomal pellets and could be dissociated by
conditions that keep the ribosomal subunits intact (8, 26, 27). Based
on this, we believe our transient expression system validly models
functional FMRP- mRNA interactions.
Although providing an important indicator, the transient transfection
data alone did not conclusively demonstrate that FMRP negatively
regulates EF-1A expression. For example, if hFMRP positively regulated
rEF-1A mRNA translation, but the hFMR1 mRNA transcription rate
exceeded its translation rate, the excess hFMR1 mRNA produced might
sequester the entire endogenous rFMRP and recombinant hFMRP. Because
the Kd values of hFMR1 and EF-1A mRNA are
similar, this would lead to decreased rEF-1A when hFMRP is expressed.
However, if the hFMR1 mRNA transcription rate is less than or equal
to its translation rate, then hFMR1 mRNA should not alter FMRP
levels enough to affect rEF-1A mRNA binding; in this case,
decreased rEF-1A would imply a negative regulatory mechanism. To
differentiate between these two possibilities, we compared EF-1A levels
in fragile X lymphoblastoid cell lines, lacking FMRP, with their normal
counterparts. We found that EF-1A protein levels were elevated in
fragile X-derived cells compared with the normal controls, whereas
EF-1A mRNA levels did not change. Both data sets are consistent
with the hypothesis that FMRP negatively regulates EF-1A expression.
Our results show that in PC12 cells, COS-7 cells, and in fragile X
lymphoblastoid cells EF-1A expression is altered as a function of hFMRP
expression. However, this observation must be extended to the brain and
testes, the major affected organs in fragile X syndrome. We are
currently addressing this by comparing EF-1A expression profiles in
fragile X knockout mouse and normal littermate controls; however,
several factors may complicate this analysis. First, it has been noted
previously that FMRP in vitro binding displays both species
specificity and isoform specificity (19), and it is currently not known
whether any mouse FMRP isoform binds mEF-1A mRNA. Second, FMRP
knockout mice demonstrate region-specific deficits in several proteins
including the receptor GluR1 (47), and their effect on EF-1A expression
is also unknown. Third, in humans, rats, and mice two forms of EF-1A
(EF-1A and EF-1A-2) are expressed in a tissue-specific manner. Unlike
lymphocytes where EF-1A is singularly expressed, both forms are
coexpressed in brain, and they appear to be regulated differentially
(48-51). Although the amino acid homology of the two proteins is high
in mice (92.4%), the nucleotide homology is less striking (79.8%), and it is not known whether EF-1A-2 mRNA binds to FMRP, or whether changes in the level of either protein may be compensated by the other.
What role might elevated EF-1A levels play in fragile X syndrome? In
yeast (52) increased EF-1A protein levels correlate with increased
nonsense suppression. In fact, a 2-fold EF-1A increase significantly
increased suppression of a number of marker genes. Such an increase is
about what we observed in fragile X lymphoblastoid cells. Whether
suppression occurs in mammalian cells in response to FMRP-altered EF-1A
expression is unknown; however, such a result might lead to increased
mutant or truncated protein levels that could negatively affect normal
cellular functions. As this would be a stochastic process this result
could explain the observed variability in the fragile X phenotype (53).
Alternatively, increased EF-1A expression might manifest itself in
growth defects and changes in cellular morphology by altering
cytoskeletal actin as has been observed in yeast (54). Actin-associated
alterations in dendritic spine shape and stability are a well known
feature of fragile X syndrome (55-57). Both questions are under active investigation. However, it is also possible that the altered EF-1A levels in fragile X patient lymphoblastoid cells have nothing to do
with brain dysfunction or any of the other clinical features of the
fragile X phenotype. Rather, it may simply be a silent by-product of
the loss of FMRP. Further work will determine which of these scenarios
is correct.