A 3′ untranslated region variant in FMR1 eliminatesneuronal activity-dependent translation of FMRP bydisrupting binding of the RNA-binding protein HuRJoshua A. Suhla, Ravi S. Muddashettyb, Bart R. Andersona,b, Marius F. Ifrimb, Jeannie Visootsaka, Gary J. Bassellb,and Stephen T. Warrena,c,d,1

aDepartment of Human Genetics, Emory University, Atlanta, GA 30322; bDepartment of Cell Biology, Emory University, Atlanta, GA 30322; cDepartment ofBiochemistry, Emory University, Atlanta, GA 30322; and dDepartment of Pediatrics, Emory University, Atlanta, GA 30322

Contributed by Stephen T. Warren, October 14, 2015 (sent for review September 29, 2015; reviewed by Claudia Bagni and Kimberly M. Huber)

Fragile X syndrome is a common cause of intellectual disability andautism spectrum disorder. The gene underlying the disorder, fragileX mental retardation 1 (FMR1), is silenced in most cases by a CGG-repeat expansion mutation in the 5′ untranslated region (UTR). Re-cently, we identified a variant located in the 3′UTR of FMR1 enrichedamong developmentally delayed males with normal repeat lengths.A patient-derived cell line revealed reduced levels of endogenousfragile X mental retardation protein (FMRP), and a reporter contain-ing a patient 3′UTR caused a decrease in expression. A control re-porter expressed in cultured mouse cortical neurons showed anexpected increase following synaptic stimulation that was absentwhen expressing the patient reporter, suggesting an impaired re-sponse to neuronal activity. Mobility-shift assays using a controlRNA detected an RNA–protein interaction that is lost with the pa-tient RNA, and HuR was subsequently identified as an associatedprotein. Cross-linking immunoprecipitation experiments identifiedthe locus as an in vivo target of HuR, supporting our in vitro find-ings. These data suggest that the disrupted interaction of HuR im-pairs activity-dependent translation of FMRP, which may hindersynaptic plasticity in a clinically significant fashion.

fragile X syndrome | FMR1 | FMRP | HuR | autism

Fragile X syndrome (FXS) is one of the most common forms ofinherited intellectual disability, and represents a well-known

genetic cause of autism spectrum disorder. FXS is a monogenicdisorder characterized by the loss or dysfunction of the fragile Xmental retardation protein (FMRP), the product of the fragile Xmental retardation 1 (FMR1) gene (1). In a vast majority of FXSpatients, FMRP expression is absent due to the expansion of anunstable CGG repeat in the promoter region of the FMR1 gene.This repeat tract is polymorphic in the population, where 5–45CGG repeats are typical. FXS patients have in excess of 200repeats, referred to as the full mutation (2), usually inherited viaan unstable maternal premutation allele (55–200 repeats). At thefull mutation length threshold of ∼200 repeats, an epigeneticevent manifests that results in hypermethylation of the FMR1promoter region and subsequent silencing of the transcript andprotein expression (3–5).Studies of FMRP function suggest that it is a selective RNA-

binding protein (RBP) that primarily acts as a negative regulator oftranslation (6, 7), and is estimated to associate with about 4–5% ofmRNAmessages expressed in the brain, including its own transcript(8–10). FMRP is also a key regulator of translation downstream ofglutamate receptor-mediated signaling in neurons, where it is rap-idly inactivated by dephosphorylation upon receptor activation,thereby allowing protein synthesis of its targets to occur in responseto the stimulus (11–14). The absence of FMRP uncouples gluta-mate receptor stimulation from the protein synthesis typically re-quired for proper signal transduction at the synapse (15). Thesemolecular defects are associated with impaired synaptic plasticity,widely believed to underlie the processes of learning and memory,which requires tightly controlled synaptic protein synthesis (16, 17)

and is thought to be a principal cause of the cognitive disabilities inFXS patients (12, 18, 19). Whereas the effects of lacking FMRPentirely have been extensively investigated, few studies have focusedon the consequences of still-present but dysregulated translationof FMRP.Despite significant analysis of the FMR1 gene, only a small

number of conventional genetic mutations, such as point muta-tions and insertions/deletions, have been reported to be associ-ated with FXS or developmental delay (20–27). To identifycauses of developmental delay attributable to FMR1 variantsother than the repeat expansion, our group sequenced the FMR1gene in 963 developmentally delayed males, each of whom testednegative for the CGG expansion mutation, and discovered a numberof previously unreported variants (28). However, the molecularconsequences, if any, of most of these variants remain unknown.In this study, we describe the functional impact of a variant

in the 3′UTR of FMR1 (c.*746T>C) using genetic, biochemi-cal, and cell biological approaches. The variant is associatedwith reduced basal FMRP levels and impairs the normal re-sponse to activity-dependent synaptic translation in culturedprimary neurons. Our data suggest that the RNA-bindingprotein HuR binds the locus normally but that this association islost when the variant is present, leading to destabilized andrapidly degraded FMR1 transcript. These findings indicate thatthe c.*746C variant allele, detected at a frequency of 1 in 160

Significance

The fragile X mental retardation protein (FMRP) is most highlyexpressed in neurons, and is critical for proper synaptic function-ing. Fragile X syndrome, a common cause of intellectual disability,is the result of absent or dysfunctional FMRP, highlighting itsimportance to the processes underlying learning and memory. Arapid upregulation of FMRP synthesis at the synapse in responseto specific neuronal signals is a key step in maintaining a dynamicsynapse, although the mechanisms governing this up-regulationare not well-understood. We show that a variant in the 3′UTRof fragile X mental retardation 1 (FMR1) causes the loss of thischaracteristic increase in synaptic FMRP synthesis, which maylead to developmental delay in patients. These data identifyseveral mechanisms and molecules modulating activity-de-pendent translation of FMRP.

Author contributions: J.A.S., B.R.A., M.F.I., G.J.B., and S.T.W. designed research; J.A.S., R.S.M.,B.R.A., and M.F.I. performed research; R.S.M. and J.V. contributed new reagents/analytictools; J.V. provided clinical support; J.A.S., B.R.A., G.J.B., and S.T.W. analyzed data; andJ.A.S., G.J.B., and S.T.W. wrote the paper.

Reviewers: C.B., Catholic University of Leuven Medical School; and K.M.H., University ofTexas Southwestern Medical Center.

The authors declare no conflict of interest.1To whom correspondence should be addressed. Email: swarren@emory.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1514260112/-/DCSupplemental.

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developmentally delayed male patients tested (28), may repre-sent an unrecognized genetic basis of developmental delay viaFMR1 dysregulation.

ResultsClinical Assessment of a Patient with the c.*746T>C Variant. One ofthe patients identified by Collins et al. (28) as harboring thec.*746T>C variant was clinically evaluated by the Emory UniversityMedical Genetics Clinic. The patient (pictured in Fig. 1A) was∼10.5 y old at evaluation and was born full-term at 10 lb, 1 ozwithout complication. He was reported to have sat independentlyat 17 mo [typically achieved by 9 mo of age (29)] and first walkedat 24 mo [typically achieved by 18 mo of age (29)]. At examination,he was nonverbal and exhibited stereotypic behavior consisting ofrocking, spinning, rubbing his fingers, and repetitively touching hisshirt collar. The patient had previously been diagnosed with au-tism spectrum disorder and attention deficit hyperactivity disorder(ADHD), and he attends special education classes. Cognitive abil-ities were assessed using the Stanford–Binet Intelligence Scales(5th Ed) (30), revealing moderate intellectual disability (IQ score47). In terms of his growth parameters, the patient is within the 50–75th percentiles for weight (88 lb), height (60 in), and head cir-cumference (55 cm). A physical examination was performedwhere bilateral flat feet with inversion were noted but no othersignificant findings. The patient’s half-brother was also evaluated,and genotyping revealed that he also possesses the c.*746C variant.He was reported to have sat independently at 17 mo old, taken hisfirst steps at 24 mo, and spoken his first word at 5 y of age. Ap-proximately 17 y old at examination, he speaks in short sentences,attends some regular 10th-grade classes with specialized classes forreading and math, is mildly intellectually disabled (IQ score 67), andhas previously been diagnosed with ADHD.

Patient 3′UTR Reduces Translation of FMRP and a Reporter. In ad-dition to the developmental delay and intellectual disability in thepatient, several noteworthy lines of evidence led us to investigatethe molecular impact of the FMR1 c.*746C allele. First, the var-iant was significantly enriched in unrelated developmentally delayedmale patients compared with gender-matched controls [found in 6of 963 patients and 0 of 1,260 controls; P = 0.007 (28)]. Addition-ally, the c.*746 position and broader genetic element are highly

conserved evolutionarily, as indicated by PhyloP (31) (2.76), GERP(32) (5.52), and PhastCons (33) (1.00) scores. Last, the locus isthymine- and uridine-rich at the RNA level, which we hypothe-sized could serve as a site of interaction for a class of U-rich RBPs(Fig. S1). Based on these data, we explored whether the varianthad any effect on FMRP expression in the patient. A lympho-blastoid cell line was established from the patient’s blood andWestern blot analysis revealed a modest, but significant, reductionof the patient’s endogenous FMRP level compared with twohealthy lymphoblastoid control lines (Fig. 1B). To corroboratethese results, luciferase reporter vectors were constructed to in-clude the full-length FMR1 3′UTR of the patient or a healthycontrol downstream of the firefly luciferase gene. We observed asignificant decrease in normalized luciferase signal with the patientreporter compared with the control in two different cell lines (P =0.007 in HEK293FT; P = 0.004 in Neuro2a; Fig. 1C). Additionally,the 3′UTR from the patient’s affected half-sibling brother, whoharbored the c.*746C allele as well, showed a similar reduction inluciferase activity compared with the control (P < 0.001; Fig. 1D).Steady-state levels of luciferase transcript were equivalent betweenthe patient and control vectors (Fig. S2), suggesting a posttran-scriptional mechanism underlying the reduction in patient reporteractivity. However, other mechanisms, such as mRNA instability,may be responsible for the observed decrease in luciferase ac-tivity, which these quantitative (q)RT-PCR assays cannot evalu-ate. Together, these data indicate that the patient 3′UTR isassociated with a decrease in endogenous FMRP and reporter ex-pression, and that the FMR1 gene may be regulated posttranscrip-tionally via the 3′UTR at the c.*746 locus.

The c.*746 Locus Is Important for Translational Regulation. To identifywhether the c.*746 locus is the specific site responsible for theobserved decrease in reporter activity because the 3′UTR ofFMR1 exhibits frequent sequence variation, we used site-directedmutagenesis to change the variant 746C nucleotide in the patientreporter to the reference thymine. This single-nucleotide changerestored luciferase activity to the level of the control. Conversely,mutagenesis of the control vector to the patient allele caused areduction in reporter signal, indicating that the variant is specificin causing the diminished translation (Fig. 2A). Given that thevariant does not completely abolish reporter activity, we examined

Fig. 1. Patient 3′UTR is associated with a significant reduction in endogenous FMRP and reporter activity in multiple cell types. (A) Photograph of a patientharboring the FMR1 c.*746C allele. (B) (Top) Representative Western blot of FMRP and eIF4e from two unrelated control male lymphoblastoid cell lines andthe patient-derived lymphoblastoid cell line. (Bottom) The band density on the Western blot of three independent protein preparations was digitallyquantified by ImageJ software (NIH). Data shown are the mean ± SD. (C) Luciferase assay in HEK293 and Neuro2a cell lines using vectors with a control 3′UTRor the patient 3′UTR. Each of three independent experiments was normalized to cotransfected Renilla luciferase activity; data shown are the mean ± SD.(D) Luciferase assay results of three independent experiments in HEK293 cells using vectors of the control, patient, and the patient’s half-brother. Data shownare the mean ± SD. Unpaired two-tailed t test, *P < 0.05; a.u., arbitrary units; n.s., not significant.

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whether mutagenesis of the broader U-rich motif would cause amore pronounced effect to determine the magnitude of trans-lational misregulation caused directly by the c.*746C variant. Wedeleted 17 bp from the locus, including the variant site, in both thecontrol and patient vectors (Fig. 2B). When expressing the controlvector with the deletion, we observed a significant decrease in re-porter activity compared with the control (P = 0.004) but not amore severe reduction than that observed with the patient reporter.The same deletion in the patient vector did not significantly alterexpression from the unchanged patient reporter (P = 0.27). Becausethe patient vector with the deletion did not show significantly al-tered expression compared with the unchanged patient vector, thec.*746 position itself is likely a critical nucleotide in the motif. Takentogether, these data implicate the locus, and the c.*746C variant inparticular, as causative of the diminished reporter expression.

The c.*746C Variant Allele Is Refractory to Glutamate Receptor Signalingin Primary Neurons. Although the patient’s lymphoblastoid cell lineshowed reduced levels of FMRP, ∼80% of normal levels were stillpresent. We questioned whether this minimal reduction in FMRPcould cause the developmental and cognitive disabilities displayedby the patient, and wondered whether there could be a neuron-specific mechanism of dysfunction that may not be apparent inother tissues. To explore this, we tested whether the variant alleleaffected activity-dependent protein synthesis in primary neurons.C57BL/6 (WT) and Fmr1 knockout (KO) mouse cortical neuronswere isolated, cultured, and transfected with the described lu-ciferase reporters to evaluate translation when treated with (RS)-

3,5-dihydroxyphenylglycine (DHPG), a glutamate analog thatstimulates synaptic activity via the group I metabotropic glutamatereceptors (mGluRs) (12, 34, 35). The addition of DHPG to cul-tures expressing the control reporter caused an increase in trans-lation in WT neurons, but not in Fmr1 KO neurons, as expected(Fig. 2 C and D, respectively). However, when expressing thepatient reporter in WT neurons, there was no increase in trans-lation after DHPG treatment in addition to the steady-state re-duction (Fig. 2C). When the mutagenized control reporter wasexpressed in WT neurons, the normal up-regulation of translationelicited by DHPG treatment was lost (Fig. 2C). These data reveala loss of activity-dependent translation in neurons in response toglutamate signaling by the patient reporter, and identify the c.*746locus as associated with this deficit.

Loss of RNA-Binding Protein Association Caused by the c.*746C Variant.Several potential mechanisms underlying these data were consid-ered, including disruption of microRNA or RBP interactionscaused by the variant allele. To determine whether an RBP targetsthe locus, we used electromobility-shift assays (EMSAs) to evaluatewhether a 42-nt biotinylated RNA probe encompassing the c.*746site interacts with a protein or protein complex from mouse whole-brain lysate. We detected a band shift using the control probe thatwas dosage-sensitive and -specific, because an excess amount ofunlabeled competitor probe was able to outcompete the labeledprobe for the interaction (Fig. 3 A and B; see also Fig. S3 and TableS1). To determine whether the c.*746C variant disrupted this in-teraction, we generated a biotinylated patient probe that differed

Fig. 2. FMR1 c.*746 site modulates steady-state and activity-dependent reporter activity. (A) Results of three independent luciferase assays using muta-genized vectors in HEK293 cells. Reference allele at c.*746, T; patient allele, C. Data shown are the mean ± SD. (B) A 17-bp deletion in the vectors that deletesthe U-rich locus including the 746 site (indicated by a horizontal bracket and red lettering, respectively) was used in luciferase assays. Results are the mean ofthree independent experiments ± SD. (C) Luciferase assay in WT mouse primary cortical neurons (E17.5) using various vectors with and without DHPGtreatment (100 μM, 5 min) ∼24 h posttransfection. Data shown are the mean of four or five independent experiments ± SD. (D) Luciferase assay in Fmr1 KOmouse primary cortical neurons (E17.5) using various vectors with and without DHPG treatment (100 μM, 5 min) ∼24 h posttransfection. Data shown are themean of four or five independent experiments ± SD. Unpaired two-tailed t test, *P < 0.05. C, control; Del, deletion; P, patient.

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only at the 746 position and compared this with the control probebinding. The patient probe lacked the band shift, suggesting a loss ofassociation with the protein or protein complex due to the 746Cvariant (Fig. 3C and Table S1). Additionally, an unlabeled com-petitor patient probe could not compete away the interaction fromthe control, even at concentrations of 40× molar excess, confirmingthe inability of the protein to bind the patient sequence (Fig. 3D).These results demonstrate that the locus is a target of an RBP orprotein complex and that the variant disrupts this association,identifying a potential mechanism underlying the observed changesin the reporter assays.

The RNA-Binding Protein HuR Associates with the c.*746 Locus. Toidentify the protein(s) that interacts with this RNA sequence, weused two approaches, both of which made use of tandem massspectrometry (MS-MS) for protein identification. First, we per-formed two biological replicates of a coimmunoprecipitation (co-IP) assay where streptavidin-coated magnetic beads were fixedwith the biotinylated control RNA probe and then incubatedwith mouse whole-brain lysate. We eluted bound proteins andsubjected the sample to MS-MS peptide sequencing to identifyenriched proteins compared with a bead-only control. Second,we performed the EMSAs as described and then excised theshifted band region from the gel and subjected the eluted proteinsample to MS-MS analysis to directly identify proteins in theband-shift region (Table S2). Three candidates were detected byboth screening methods: HuR (also known as ELAVL1), PUR-α,and PUR-β. Whereas the PUR proteins typically bind purine-richmotifs, HuR is known to target U-rich elements in the 3′UTR ofgenes. Given the U-rich nature and location of the FMR1 c.*746locus, we pursued HuR as the top RBP candidate.To determine whether the band shift we observed in the EMSAs

was indeed HuR, we first performed a gel-supershift assay using anantibody to HuR. As shown in Fig. 4A, the shifted band disappearswith the addition of HuR antibody to the binding reaction, althoughthe supershifted band was not visible due to the high backgroundsignal inherent to using a whole-brain lysate. Next, we used a HuRantibody to immunopurify HuR-bound mRNAs from HEK293 celllysates and then assayed for FMR1 as well as an established targetof HuR (β-actin) and a gene not known to interact with HuR(GAPDH). FMR1 was enriched by HuR IP ∼400-fold over an IgG

control, similar to the level of β-actin enrichment, whereas GAPDHshowed no enrichment (Fig. 4B), indicating that HuR indeed as-sociates with FMR1. To localize this interaction specifically to the746 locus, we performed gel-shift, competition, and supershift as-says with purified HuR protein rather than whole-brain lysate.These assays revealed an association between HuR and the controlprobe, and confirmed the lack of interaction between the patientprobe and HuR (Fig. 4C). Altogether, FMR1 is bound by HuRat the c.*746 locus, and this interaction is disrupted by thevariant allele.

HuR Localizes to Dendrites and Synapses. HuR is typically mostabundant in the nucleus of cells, but is known to translocate tothe cytoplasm under certain conditions to facilitate mRNA sta-bility and translation (36, 37). If HuR is involved in the activity-dependent translation of FMR1, it must be present at the syn-apse, because previous work has shown that endogenous FMR1mRNA localizes to dendrites (38) and can be locally translated inan activity-dependent manner (39). To determine whether HuRis present at the synapse, we used immunocytochemistry to vi-sualize its location in primary cortical neurons. Although themajority of HuR is located in the nucleus, it is also present in thedendrites distal to the soma (Fig. 4D), a finding supported byother very recent data (40). To more specifically determine whetherHuR localizes to the synapse, we performed immunoblot analysesfrom mouse synaptoneurosome preparations and found the proteinto be present, suggesting that HuR is at the synapse (Fig. 4E).Previous work has shown that Fmr1 mRNA is present in synapticfractions (18), so we investigated whether Fmr1 mRNA was in acomplex with HuR in this compartment. Co-IP experiments insynaptoneurosomes revealed a significant enrichment of Fmr1mRNA in HuR IP fractions, suggesting that these molecules in-teract at the synapse (Fig. 4F).

Patient FMR1 mRNA Decays Rapidly but Is Trafficked to DendritesNormally. HuR serves many functions, including translationregulation, transcript trafficking, and splicing. One of the best-studied functions of HuR is that of mRNA stabilization oftranscripts that it binds via U-rich elements in the 3′UTR (36).Because HuR binding to the patient sequence appears to beimpaired, we assessed the stability of endogenous FMR1 mRNAin the patient lymphoblastoid cell line. We performed mRNA

Fig. 3. RNA EMSAs reveal a specific and dose-dependent interaction between a protein and the c.*746 locus that is absent with the patient sequence. (A) Abiotinylated control RNA probe incubated with increasing amounts of mouse whole-brain lysate and resolved by 5% native TBE gel. (B) An unlabeled versionof the control RNA probe was added in increasing amounts (up to 40× excess by molar concentration) to the whole-brain lysate binding reaction and resolvedon a 5% native TBE gel. (C) A biotinylated patient RNA probe containing the c.*746C nucleotide was incubated with increasing amounts of mouse whole-brain lysate in the same manner as the control probe and resolved on a 5% native TBE gel. (D) An unlabeled version of the patient RNA probe was added inincreasing amounts, up to 40× molar excess concentrations, to the binding reactions with the labeled control probe and resolved on a 5% native TBE gel.

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Fig. 4. HuR binds FMR1 at the c.*746 locus and is present at the synapse. (A) Supershift assays were performed using 32P-labeled control probe and HuRantibody. Binding reactions were carried out as described previously with the addition of 0.5–1 μg of antibody per 10 μg of mouse whole-brain lysate ineach reaction. (B) Immunoprecipitation of HuR protein from HEK293 cell lysate was performed, and copurified mRNAs were assayed by real-time RT-PCR.The results show the fold enrichment of mRNA levels of β-actin (a known target of HuR), GAPDH (not known to interact with HuR), and FMR1 mRNAnormalized to IP with IgG alone. All data are displayed as the ratio of IP mRNA levels to input mRNA levels. Data shows the mean of three independentexperiments ± SD. (C) RNA EMSA using purified HuR protein with the control or patient probe, unlabeled competitor control probe, and HuR antibody.(Bottom) Free probe bands from a shorter exposure of the same blot to ensure equal probe loading. (D) Image of immunofluorescence staining for HuR(green), PSD-95 (red), and nuclei (blue) in primary cortical neurons. A merged image of all colors is shown (Top Left). (Scale bar, 10 μm.) The white boxes inthe merged image are enlarged in the Bottom two rows, with the Top row representing the Left box and the Bottom row representing the Right box. Themerged, HuR (green), and PSD-95 (red) images are shown in both Bottom rows from Left to Right, respectively. (Scale bars, 5 μm.) (E ) Western blotshowing the presence of HuR in a whole-cortical lysate and synaptoneurosomes. GFAP, a glial marker, was used as an indicator of synaptoneurosomepurity. An antibody against eiF4e was used as a loading control. C1 and S1, cortical lysate and synaptoneurosomes, respectively, of preparation 1; C2 andS2, cortical lysate and synaptoneurosomes, respectively, of preparation 2. (F ) Synaptoneurosome preparations were subjected to HuR immunoprecipi-tation, and the level of associated Fmr1 was assayed by qRT-PCR. The levels of GAPDH and β-actin mRNA were also assayed in the immunoprecipitate as anegative and positive control, respectively. Data shown are the mean of three independent synaptoneurosome preparations ± SD. (*P < 0.05 using a two-tailed unpaired t test.) (G) Actinomycin D (5 μg/mL) was added to control and patient lymphoblastoid cells to block transcription, and qRT-PCR wasperformed for FMR1 (Right) and PSD-95 (Left) after 1, 2, 4, and 6 h of drug treatment as well as an untreated control (0 h). Each point shows the meanpercentage of mRNA remaining relative to the 0-h control ± SEM in six independent experiments. Nonlinear regression lines were fit to each set of datapoints, and the slopes were calculated and compared. For FMR1, the difference in slope was significantly different between the control and patient (P =0.008); for PSD-95, the difference in slope was not significantly different (P = 0.582).

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decay assays using actinomycin D to block transcription and thentracked the longevity of the extant transcript over time. Weobserved rapid decay of FMR1 in the patient cell line, which wassignificantly faster than a control lymphoblastoid cell line (P =0.008). Another gene, PSD-95, showed no difference in the rateof mRNA degradation after actinomycin D treatment betweenthe patient and control (P = 0.582; Fig. 4G). These results suggestthat the stability of endogenous FMR1 in the patient is diminished,potentially because of the inability of HuR to bind at the c.*746locus and stabilize the transcript. This finding may explain thesteady-state reduction in reporter assays and endogenous FMRPin the patient cell line, and potentially has implications for theactivity-dependent defect observed in cultured neurons.Another known function of HuR is trafficking of its mRNA

targets to appropriate cellular compartments (41). Because HuRtargets FMR1, we sought to determine whether the patient tran-script was being properly transported to locations distal to thesoma, as this may be controlled in part by HuR and may explainthe defect in activity-dependent translation of the reporter incultured neurons. Here we used fluorescently tagged FMR1 totrack and compare the mutant transcript shuttling and localizationin primary neurons compared with a control. Both versions oftagged transcript were observed in dendrites and colocalized withHuR and FMRP, suggesting that the patient message is beingtrafficked to the appropriate neuronal compartments (Fig. S4).We also tracked the movement of the FMR1mRNAmessage overtime to determine whether the rate of localization of the patienttranscript was impaired relative to the control, and did not observeany significant difference (Fig. S5). Thus, the c.*746 locus likelydoes not play a role in FMR1 trafficking.

HuR Targets the c.*746 Locus in Vivo, and Knockdown of HuR ReducesFMR1 and FMRP Levels. We next wanted to determine whether theHuR interaction at the c.*746 locus is recapitulated in vivo andwhether disruption of the interaction has an effect on FMR1 andFMRP expression. To do this, we analyzed HuR binding profilesgenerated in three published datasets: two that used photo-activatable-ribonucleoside–enhanced cross-linking immunoprecipita-tion (PAR-CLIP) (42, 43) and one study comparing multiple CLIPprotocols (44). CLIP assays capture in vivo interactions by UV-irra-diating live cells or tissue at a wavelength that cross-links the RBP tothe target RNA, thereby identifying specific sites or motifs of contact(45–47). All three studies identified the 3′UTR of FMR1 as a targetof HuR. Although the sites of FMR1–HuR interaction differ slightlybetween the datasets, the Kishore et al. study (44) detected HuRassociation at the c.*746 locus through two different CLIP protocols,linking the patient variant site to HuR binding in vivo. The other twostudies found HuR binding sites near the c.*746 locus, although theCLIP sequence tags do not directly overlap with the c.*746 site(Table S3). Taken together, there is evidence that the c.*746 site istargeted by HuR in vivo, although more studies will be necessary toconfirm these findings.To examine the effect of HuR on its target genes, Lebedeva et al.

(42) measured new protein synthesis and changes in mRNA levelsusing pulsed stable isotope labeling by amino acids in cell culture andRNA-sequencing experiments, respectively, in cells depleted of HuRand calculated the log2 fold change compared with a negative con-trol for each target gene. Using these data, we plotted the distribu-tion of changes in mRNA levels after HuR knockdown anddiscovered that FMR1 levels were significantly reduced comparedwith other genes assayed (log2 score −0.87; Fig. 5A). Additionally,FMRP was the third–least-abundant protein in the absence of HuRfrom over 2,000 proteins analyzed (log2 score −1.37; Fig. 5B). Evenwhen normalizing protein synthesis by concomitant changes inmRNA levels, FMRP was still among the least-synthesized proteins(log2 score −0.51; Fig. 5B), indicating a role for HuR in translationregulation of FMR1. These data suggest that FMR1 is targeted by

HuR at the c.*746 locus in vivo and is positively regulated by theprotein, consistent with our in vitro results.

DiscussionThe variant c.*746T>C in the FMR1 3′UTR reduces the basallevel of FMRP in patient-derived cell lines and in multiple othercell types using reporter constructs and, perhaps more importantly,eliminates the normal response to glutamate signaling by reporterassay in primary cultured neurons. Our data suggest that bothdefects are the consequence of a disrupted RNA–protein in-teraction between HuR and FMR1 caused by the c.*746C variant.HuR is known to bind U-rich motifs in 3′UTRs and introns,particularly stretches of uridines with interspersed adenines orguanines (42, 43, 48). The change from uridine to cytosine at thec.*746 position, which interrupts a multiuridine stretch, presumablyhinders the normal interaction of HuR at the locus. In support ofthis hypothesis, mutagenesis of the variant locus modulated thesteady-state expression of the patient and control vectors as well asthe activity-dependent translation. The c.*746C patient allele resultedin a molecular phenotype similar to that observed in Fmr1 KOneurons after glutamate receptor activation, where glutamate-drivenprotein synthesis was absent. Although these molecular findingsare compelling, they do not definitively link the c.*746C variantto the patient’s phenotype. Consequently, it will be important toevaluate more families and individuals with the c.*746C allele todetermine whether the variant is truly pathological and observethe range of severity and disabilities caused by the variant. Thepatient’s half-brother is also developmentally delayed/intellectuallydisabled, and his FMR1 3′UTR showed a reduction in expression byreporter assay, which provides a second case (albeit familial) thatmay be attributable to the c.*746C allele. However, he is affected toa lesser degree compared with the proband, which may indicate thatother genetic or environmental factors specific to the proband playa role in the relative severity of the phenotype.The role of synaptically expressed FMRP is hypothesized to

function as a negative feedback loop, where existing FMRP isdephosphorylated and degraded after glutamate receptor stimula-tion and newly synthesized FMRP reins in the burst of translationafter an appropriate amount of time (14, 34, 49). However, theimportance of locally translated FMRP had not been directlyaddressed until recently, when it was discovered that a premutationFXS mouse model exhibited impaired activity-dependent FMRPsynthesis likely due to the presence of expanded CGG repeats inthe 5′UTR (50). Iliff et al. (50) exploited a well-characterizedneuronal phenotype in Fmr1 KO mice, enhanced mGluR-medi-ated long-term depression (LTD), to evaluate the impact of dys-functional local FMRP translation on synaptic plasticity inpremutation brain tissue. Their results suggest that the reducedlevels of pre-existing FMRP were not sufficient to suppress theenhanced mGluR-LTD occurring in the premutation neurons; lo-cally synthesized FMRP was necessary for proper LTD. Whereasmost studies of the premutation allele have focused on late-onsetneurodegenerative phenotypes, accumulating evidence suggeststhat the premutation may have a demonstrable effect on neuro-development as well (51–53). Although the genetic defect inmGluR-mediated translation studied by Iliff et al. (i.e., the pre-mutation in the 5′UTR) differs from the 3′UTR defect investigatedhere, it suggests that the c.*746C variant could hinder neuro-development and synaptic plasticity through the impairment of ac-tivity-dependent translation, and is a possible reason for the observeddevelopmental delay and cognitive disability in the patient.These data implicate the c.*746 locus in the basal and activity-

dependent expression of FMRP. However, the exact mechanism(s)by which this deficit occurs remains to be determined. The trans-lational insufficiencies could be a direct effect of the inability ofHuR to bind FMR1 or an indirect consequence of this perturbedinteraction, such as rapidly decaying transcript at the synapse that isinsufficient to support activity-regulated protein synthesis. Another

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potential mechanism is based on the observation that HuR com-petes with other binding proteins, of which there are several thatrecognize a similar U-rich motif (36), to modulate the stability of agiven target transcript. For example, the binding factor AUF1 (alsoknown as hnRNP D), which was also pulled down in our co-IP/MSscreen (Table S2), is associated with rapid degradation of boundmRNAs, opposing the function of HuR. Indeed, this type ofcompetitive binding has recently been postulated to occur in vivobetween HuR and another U-rich binding protein that negativelyregulates transcript stability, ZFP36 (54). This presents the possi-bility that each is required as an antagonist to the other, and eachperforms their specific functions when cued by the appropriatecellular signals. If HuR binding is impaired at the c.*746 site,AUF1 or other similar transcript-destabilizing proteins may havean increased opportunity to degrade FMR1, leading to a reducedmRNA half-life, which we observed in the patient cell line.A recently recognized function of HuR is that it can act as an

“anti-RISC” (RNA-induced silencing complex); that is, HuRbinds near microRNA (miRNA) sites, oligomerizes along themRNA, and removes or blocks the miRNA machinery fromreaching its target, thereby relieving the down-regulation typicallyimposed by miRNAs (43, 55, 56). One study has shown that miR-130b, an miRNA that is expressed in the brain and is highlyconserved, interacts at the 746 variant locus (seed sequence bindsat FMR1 c.*755–761), negatively regulates the expression ofFMR1 in neural progenitor cells, and affects cell-fate specification(57). These findings present an alternate mode of dysfunctionwhereby the inability of HuR to bind and/or oligomerize adjacentto the miR-130b site hinders the removal of the miRNA, whichthen continues to repress FMRP translation unchecked and per-haps alters the response to mGluR stimulation. There is evidenceof an miRNA-mediated response to glutamate signaling, whichrequires the removal of miR-125a to allow the normal increase inPSD-95 expression following glutamate receptor activation (58).If miR-130b is involved in activity-dependent translation ofFMRP, failure to block or eliminate its association with FMR1mayprevent the typical up-regulation of FMRP synthesis followingglutamate signaling. Although the disrupted FMR1–HuR interac-tion may be the basis of the observed dysfunctions, more researchis required to identify the process(es) that is perturbed.Multiple different approaches in this study demonstrate that

HuR binds the FMR1 c.*746 locus in vitro. To investigate in vivobinding of HuR to the locus, we analyzed three published datasetsthat identified mRNA targets and binding locations of HuR byPAR-CLIP assays in human cell lines (HEK293 and HeLa) (42–44). All datasets showed that multiple 3′UTR and intronic sites ofFMR1 are targeted by HuR. Because the specific binding locations

identified by each dataset were inconsistent, it will be important toconfirm these findings with more targeted studies. Additionally, itwould be of interest to determine whether the interactions arerecapitulated in different cell types, including neuronal cells orbrain tissue. To determine the functional effect of HuR on FMR1,we analyzed data from experiments that tracked the translation ofnew proteins after knockdown of HuR, which revealed a drasticreduction in FMR1 mRNA levels and FMRP synthesis. In fact,FMRP levels were reduced more than all other proteins assayedexcept for two, one of which was HuR itself. Together, these re-sults suggest that the locus is targeted by HuR and that FMRPexpression is significantly down-regulated in the absence of HuR.If HuR-mediated regulation of FMR1 is via the 746 locus, the lackof HuR binding to the patient allele may underlie the reduction inexpression and transcript stability observed in our data.Because HuR is a member of a small protein family (HuR,

HuB, HuC, and HuD) and the other Hu paralogs also bind asimilar U-rich sequence motif (59), we were interested in whetherthese other Hu proteins associated with FMR1 at the c.*746 locus.Our co-IP followed by MS-MS assay did identify HuC as beingcopurified with the biotinylated probe, although it was not de-tected in the gel slice/MS-MS assay and, therefore, was not in-cluded in our top candidates. However, these data do suggest atleast some level of interaction between the neuronal Hu and thec.*746 locus. Additionally, the neuronal Hu proteins were foundto bind the c.*746 locus in vivo in mouse brain on actively trans-lating polyribosomes (10). Taken together, the locus is a target ofposttranscriptional regulation, and may be targeted by differentHu proteins under specific temporal, spatial, and environmentalconditions, a possibility that should be examined further.The findings presented here illustrate the impact of a single-

nucleotide variant in the regulatory region of a gene, which canhave significant molecular consequences and may be causative of aclinical phenotype. As whole-genome sequencing becomes morecommonplace, sequence data in the UTRs and other regulatoryregions will be available for analysis and should be explored forfunctional variants like the one described here when studying thegenetic defects underlying certain diseases. Together, these dataidentify the FMR1 c.*746 locus as an important site of post-transcriptional regulation and shed new light on the mechanismsgoverning activity-dependent translation of FMRP.

Materials and MethodsHuman Subject and Animal Research. All experimental procedures requiringmouse models was approved by the Emory University Institutional ReviewBoard (IRB). Informed consent was obtained from all family members de-scribed in the study and was approved by the Emory University IRB.

Fig. 5. HuR depletion down-regulates FMR1 mRNA levels and FMRP synthesis. (A) The distribution of mRNA changes after knockdown (KD) of HuR reveals asignificant decrease in FMR1 levels compared with over 2,000 other mRNAs assayed (log2 value −0.87). The red arrow indicates the bin in which FMR1 falls.(B) The distribution of protein-level changes shows a significant decrease in FMRP synthesis (log2 value −1.37). The red arrow indicates the bin in which FMRPfalls. (C) The distribution of protein synthesis changes when accounting for concomitant mRNA changes indicates that the decrease in protein synthesis is notonly the result of a reduction in mRNA levels (i.e., −log2 difference between protein and mRNA levels −0.51). The red arrow indicates the bin in which FMRPfalls. All histograms were generated using data from Lebedeva et al. (42).

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Luciferase Constructs and Site-Directed Mutagenesis. The Dual-LuciferaseSystem from Promega was used for reporter assays. In short, the control andpatient 3′UTRs were amplified with a forward primer that included a 5′ XbaIsite and a reverse primer that included a 5′ BamHI site. These amplicons werecloned using the TOPO TA Cloning System (Invitrogen), excised using BamHIand XbaI restriction enzymes, and ligated into a pGL3-promoter vectordownstream of the luciferase gene. All constructs were sequenced to verifyproper orientation and sequence. The pRL-SV40 vector was used as a trans-fection normalization control. The control and patient 3′UTR vectors were usedas templates for all mutagenesis experiments using the Agilent QuikChangeLightning Kit as recommended.

Cell Culture and Transfection. Neuro2a cells were grown in DMEM supple-mented with 10% (vol/vol) FBS and 1× penicillin/streptomycin antibiotics;HEK293FT cells were grown in DMEM supplemented with 10% FBS, 2 mML-glutamine, 1× penicillin/streptomycin, 0.1 mM nonessential amino acids, andgeneticin-selective antibiotic. Both cell lines were grown at 37 °C with 5% CO2.

Transfections in HEK293FT and Neuro2a cells were performed using Lip-ofectamine 2000 or Lipofectamine LTX (Invitrogen) in 6- or 12-well platesaccording to the manufacturer’s instructions. Lipofection complexes wereleft in the medium for 4–6 h and then removed and replaced with freshmedium. Cells were allowed to express the firefly and Renilla luciferase for∼24 h before harvesting.

Primarymouse cortical neuron [embryonic day (E)17.5] cultureswere isolatedfrom C57BL/6 and Fmr1 KO embryos and plated in 12-well plates coated with0.2 mg/mL poly–L-lysine. Transfections were carried out at 13 d in vitro usingNeuroMag Transfection Reagent (OZ Biosciences) following the recommendedprotocol. Neurons were stimulated with 100 μM (RS)-3,5-DHPG (Tocris Biosci-ences) for 5 min before lysis with passive lysis buffer (Promega).

Luciferase Assay. Lysis of the cells was performed using the passive lysis protocolfrom the Dual-Luciferase Assay Kit (Promega). To detect luciferase signal, 10–20 μLcell lysate and 50 μL Luciferase Assay Reagent II were mixed thoroughly bypipetting and placed in the luminometer, and firefly luminescence readingswere collected. Immediately after, 50 μL Stop & Glo Reagent (Promega) wasadded and mixed thoroughly by pipetting, and the Renilla luminescence signalwas detected. All values reported are the ratio of firefly:Renilla reporter signal.

Quantitative RT-PCR. Reverse-transcription reactions were carried out usingiScript One-Step RT-PCR or iTaq Universal SYBR Green Kits (Bio-Rad). All qPCRreactions were performed in duplicate or triplicate as technical replicates. Allreactions were cycled for 40 cycles using the Bio-Rad CFX96 real-time system,followed by a melt curve cycle, and then analyzed with the Bio-Rad CFX96software package.

Immunoprecipitation. Dynabeads MyOne Streptavidin T1 (Thermo FisherScientific) was used for biotinylated RNA probe IP following the manufac-turer’s recommended protocol. In each of two independent assays, 50 μg ofbiotinylated RNA probe was immobilized on 200 μL washed Dynabeads sup-plemented with cOmplete, Mini, EDTA-free (Roche) and SUPERase·In (LifeTechnologies) and then incubated with 1 mg whole-brain lysate and a 10× ex-cess of Torula yeast RNA (Sigma-Aldrich) in binding buffer (1 M NaCl, 1 mMEDTA, 10 mM Tris·HCl, pH 7.5) for 1 h at room temperature with rotation.After three washes, coimmunoprecipitated proteins were eluted in 30 μLbinding buffer with 0.1% (wt/vol) SDS at 95 °C for 5 min and subjected tomass spectrometry.

HuR antibody (MBL; RN004P) was immobilized on Dynabeads Protein G(Immunoprecipitation Kit; Life Technologies) and incubated with HEK293 celllysates at room temperature for 12 min by rotation, along with a proteinG-only control. After four washes, the coimmunoprecipitated RNA was elutedwith 50 μL elution buffer at 95 °C for 5 min and then purified with TRIzol LS(Life Technologies). qRT-PCR was performed using FMR1, β-actin, andGAPDH primers for an input sample as well as the total amount of IP sample,and the ratio of calculated starting quantity of IP:input was used to de-termine the enrichment of each mRNA target.

Western Blotting. Lysates from lymphoblast cell lines were generated byresuspending cells in lysis buffer [50 mM Tris·HCl, 300 mM NaCl, 30 mM EDTA,0.5% Triton X-100, pH 7.6; cOmplete, Mini, Protease Inhibitor Tablet (Roche)]and incubated on ice for 15min, followed by pelleting of cell debris at ~23,000×gfor 15 min at 4 °C. SDS/PAGE (4–15% gradient) was performed with 10–20 μgtotal lysate as determined by the Bradford assay. Proteins were trans-ferred to PVDF, blocked with T20 PBS blocking buffer (Thermo Scientific)for 15–30 min, and incubated with primary antibodies [anti-FMRP (Millipore;MAB2160) at 1:2,000 dilution and anti-eIF4e (BD Biosciences; 610270) at1:2,000 dilution] overnight at 4 °C with gentle agitation. Next, the membranewas incubated with HRP-conjugated anti-mouse and anti-rabbit secondaryantibodies at 1:10,000 dilution in 1% (wt/vol) milk blotto for 1 h at room tem-perature, washed three times with 1% milk blotto, and incubated with HyGLOECL solution (Denville Scientific) for 1 min with agitation.

RNA Probe Generation by in Vitro Transcription. DNA templates containing theT7 promoter and probe template sequence were generated by a modifiedoverlap extension PCR strategy (60). Each oligonucleotide (2 μM) was addedto a PCR mix consisting of 1× Herculase II reaction buffer, 0.25 mM eachdNTP, 0.5 μL Herculase II fusion enzyme (Agilent Technologies), 3 μL 5 Mbetaine solution (Sigma-Aldrich), and MilliQ water up to 50 μL. The reactionwas cycled under the following parameters: 95 °C for 2 min, followed by 35cycles of 95 °C for 20 s, 45 °C for 30 s, and 72 °C for 30 s. A final extensioncycle of 3 min at 72 °C was added. The resulting double-stranded PCRproducts were run on a 1% (wt/vol) agarose gel, extracted from the gel(Qiagen MinElute Gel Extraction Kit), and resuspended in MilliQ water. DNAtemplates were used at a concentration of 2 pM in the in vitro transcriptionreaction for 3–4 h at 37 °C (Ambion T7 MEGAshortscript Kit) in the presenceof biotin-17-ATP (Enzo Biosciences) or biotin-14-CTP (Life Technologies). Theresulting 42-nt RNA transcripts were purified using TRIzol LS (Ambion). Thesame in vitro transcription procedure was used to generate unlabeled RNAprobes for 32P-end labeling and competition gel-shift assays.

Electromobility-Shift and -Supershift Assays. Gel-shift assays were carried outusing the LightShift Chemiluminescent RNA EMSA Kit (Thermo Scientific)according to the manufacturer’s protocol. Briefly, biotin-labeled RNA probeswere mixed with C57BL/6 (WT) or FMR1 KO mouse whole-brain lysates and in-cubated at room temperature for 20–30 min and then electrophoresed througha 5% nondenaturing Tris-borate-EDTA (TBE) gel. The RNA–protein complexeswere transferred to a positively charged nylon membrane and developed to filmfollowing the RNA EMSA protocol. Supershift assays were carried out in a similarmanner with the following exceptions: addition of ∼1 μg of antibody and in-cubation at room temperature for 20 min before the addition of the radiola-beled or biotin-labeled RNA probe in a total of 20 μL reaction volume.

mRNA Decay Assay. Lymphoblastoid cells were plated in six-well plates at adensity of 5 × 106 cells per well in 2 mL medium consisting of RPMI-1640, 10%FBS, 2 mM L-glutamine, and 1× penicillin/streptomycin antibiotic. Cells weretreated with 5 μg/mL actinomycin D (Sigma) for various amounts of time andthen harvested by TRIzol LS (Ambion) extraction according to the manufacturer’srecommended protocol. qRT-PCR was used to assess the amount of FMR1 tran-script remaining compared with an untreated control. All qRT-PCRs were per-formed in duplicate, and the mean level of FMR1 remaining at each time pointwas normalized to the mean level of β-actin remaining at each time point.

ACKNOWLEDGMENTS.We thank Heather Clark for contacting the patient andrecording the family history; Duc Duong, Nicholas Seyfried, and the EmoryUniversity Proteomics Core Facility for mass spectrometry data and advice;Tamika Malone for mouse colony maintenance and cortical tissue dissections;Pankaj Chopra for rank aggregation analysis of mass spectrometry assays; MikaKinoshita, Leila Myrick, and Michael Santoro, among others in the S.T.W. andG.J.B. laboratories, for helpful insight and discussion. We also thank RobertDarnell and Jennifer Darnell for helpful discussions and data sharing. Thiswork was supported by NIH Award NS091859 from the National Instituteof Neurological Disorders and Stroke; Eunice Kennedy Shriver NationalInstitute of Child Health and Human Development in support of the EmoryNational Fragile X Research Center (S.T.W.); NIH Award 1R21NS091038 (toG.J.B.); and a FRAXA Research Foundation fellowship (to J.A.S.).

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