MOLECULAR AND CELLULAR BIOLOGY, Dec. 2002, p. 8438–8447
0270-7306/02/$04.00⫹0 DOI: 10.1128/MCB.22.24.8438–8447.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Vol. 22, No. 24

Casein Kinase II Phosphorylates the Fragile X Mental Retardation
Protein and Modulates Its Biological Properties
Mikiko C. Siomi,* Kyoko Higashijima, Akira Ishizuka, and Haruhiko Siomi
Institute for Genome Research, University of Tokushima, Kuramoto, Tokushima 770-8503, Japan
Received 28 May 2002/Returned for modification 12 August 2002/Accepted 6 September 2002

Fragile X syndrome is caused by loss of FMR1 protein expression. FMR1 binds RNA and associates with
polysomes in the cytoplasm; thus, it has been proposed to function as a regulator of gene expression at the
posttranscriptional level. Posttranslational modification of FMR1 had previously been suggested to regulate its
activity, but no experimental support for this model has been reported to date. Here we report that FMR1 in
Drosophila melanogaster (dFMR1) is phosphorylated in vivo and that the homomer formation and the RNAbinding activities of dFMR1 are modulated by phosphorylation in vitro. Identification of a protein phosphorylating dFMR1 showed it to be Drosophila casein kinase II (dCKII). dCKII directly interacts with andphosphorylates dFMR1 in vitro. The phosphorylation site in dFMR1 was identified as Ser406, which is highly
conserved among FMR1 family members from several species. Using mass spectrometry, we established that
Ser406 of dFMR1 is indeed phosphorylated in vivo. Furthermore, human FMR1 (hFMR1) is also phosphorylated in vivo, and alteration of the conserved Ser500 in hFMR1 abolishes phosphorylation by CKII in vitro.
These studies support the model that the biological functions of FMR1, such as regulation of gene expression,
are likely regulated by its phosphorylation.
individuals (18). These data suggest that FMR1 takes part innormal synaptic formation during development by regulating
the expression of specific mRNAs at the postsynaptic sites.
What kind of mRNAs does FMR1 specifically bind to in
vivo? It has been reported that FMR1 in vitro binds specificallyto poly(G) and poly(U), but not to poly(A) and poly(C) (38),and it was estimated that FMR1 interacts with about 4% of
human fetal brain mRNAs (2). These results indicate that
FMR1 has an intrinsic ability to distinguish its target moleculesfrom among all the native RNA molecules and to bind specificsequences. Recently, mRNAs to which FMR1 preferentiallybinds have been identified by several groups (5, 7, 36, 53). In
the mRNAs identified, the nucleotide sequence through which
FMR1 prefers to bind can form unusual secondary structures
known as G quartets (7, 36), which were originally reported as
DNA structures at the ends of chromosomes for stabilization(43, 51). The G quartet structure in RNA is known to bestabilized by coordination with particular cations, such as K⫹.
Indeed, FMR1 showed much higher affinity to a G quartet ina buffer containing K⫹. Interestingly, Darnell et al. reported
that only the RGG box in FMR1 is required, and that the two
KH domains are dispensable, for the interaction between
FMR1 and a G quartet (7). However, the in vivo relevance of
these discoveries has not been elucidated.
The discovery of the existence of two FMR1-related genes,
FXR1 and FXR2, has revealed an additional level of complexity
in the study of FMR1 functions in vertebrates (40, 52). For
many human disorders, redundancy of gene function due togene duplication in the same species makes it difficult to interpret clearly the effects of loss-of-function mutations on
cellular physiology and biochemistry. Recently, a Drosophila
homolog of FMR1 (dFMR1) has been identified (48). In agreement with the statement of Wan et al. (48), the recently completed Drosophila melanogaster genome sequence (1) revealed
that dFMR1 is a unique, single gene with no homologs in the fly

Fragile X syndrome is the most frequent cause of inheritablemental retardation and is also one of the most common singlegene disorders (14). Cognitive deficits reported for fragile X
children range from mild to severe, and behavioral disturbances include social and attention deficits, autistic-like behaviors, unusual responses to sensory stimuli, hyperactivity, andsleep problems (12, 14). The gene directly responsible forfragile X syndrome, FMR1, was identified in 1991 (47). In mostcases, the syndrome is caused by a trinucleotide repeat expansion in the 5⬘ untranslated region of the FMR1 gene (49). The
expansion of the CGG repeat results in an absence of theencoded protein. It is therefore clear that the pathophysiological mechanisms leading to symptoms in fragile X syndrome
can be elucidated by studying the functions of the FMR1 gene.
The fact that the FMR1 protein is a cytoplasmic protein with
RNA-binding motifs (two KH domains and an RGG box) (2,38, 39) and the fact that it associates with ribosomes (6, 9, 21,41) suggest that this protein functions in the posttranscriptional regulation of some specific mRNA expression (16). Indeed, FMR1 has recently been proposed to be a translational
repressor in a cell-free system and in frog oocytes (26, 27).
Interestingly, although the inhibitory effect was demonstrated
not to be influenced by the features of the mRNAs used inthose studies, FMR1 seemed to specifically inhibit its own
translation (36). It has been known that FMR1 is found at
postsynaptic sites enriched in ribosomes, where some specificmRNAs, including FMR1, are translated in response to synaptic activation (13). It has also been observed that in fragile X
patients the numbers of dendritic spines are different and themorphology is also changed from that of those cells in normal* Corresponding author. Mailing address: Institute for Genome Research, University of Tokushima, Kuramoto, Tokushima 770-8503,
Japan. Phone: 81-88-633-9456. Fax: 81-88-633-9451. E-mail: siomim@genome.tokushima-u.ac.jp.
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genome. The Drosophila and vertebrate FMR1 proteins sharea number of topographical landmarks, including RNA-binding
motifs (two KH domains and an RGG box), and show similar
biological properties such as a binding preference in vitro forcertain RNA sequences and ribosomal association in vivo (19,48; M. C. Siomi and H. Siomi, unpublished data). Moreover, anumber of genes related to complex behaviors such as learningand memory are conserved between flies and humans (30, 34).
Therefore, Drosophila could be used to define the molecularpathways leading to human neurological diseases such as fragile X syndrome and to identify the genes involved. In fact,
several groups have individually produced fly models of fragile
X syndrome and have shown that dFMR1 is necessary fornormal synaptic activities (53) and for rhythmic circadian output regulating locomotor activities (8, 17, 31).
For a number of proteins, posttranslational modifications
have been well studied and are known to be critical for regulating physiological and biochemical functions. So far, however, very little is known about the posttranslational modifications of FMR1 protein and their effects at the functional level.
Here we provide the first evidence that FMR1 protein, both inflies and in humans, is phosphorylated in vivo. By purificationfrom the cytoplasmic lysate of a Drosophila culture cell line,
Schneider 2 (S2), and by mass spectrometric analysis, we identified Drosophila casein kinase II (dCKII) (35) as a protein
phosphorylating dFMR1. Indeed, recombinant dCKII was able
to interact with and phosphorylate dFMR1 in vitro. Systematic
mutation analyses revealed that the phosphorylation site indFMR1 is Ser406, located just upstream of the RGG box. Thisamino acid residue is conserved among FMR1/FXR family
members across many species, and mutation to Ala of the Ser
residues in dFMR1 and hFMR1 proteins abolishes the phosphorylation by CKII in vitro. Finally, we demonstrate that theefficiency of dFMR1 in forming homomers and binding RNA
is profoundly altered by phosphorylation in vitro. Taken together, our data strongly support the hypothesis that phosphorylation by CKII could play an important role in regulating
physiological functions of FMR1, such as regulation of gene
expression for the normal formation of synapses during development.

In vitro phosphorylation assay. Either full-length or truncated dFMR1 cDNA(48) was subcloned into the pGEX-5X (Amersham Bioscience) bacterial expression vector to produce glutathione S-transferase (GST) fusion proteins for assay.
GST fusion proteins and GST itself were produced and purified as described by
the manufacturer. For Fig. 6C, full-length hFMR1 cDNA (38, 47) was used and
GST-hFMR1 was produced as well. All the point mutants were constructed by
performing PCR using primers specifically designed to alter Ser to Ala. The final
products were inserted into the pGEX-5X expression vector, and GST fusion
proteins were produced as well. The assay was carried out in P buffer containing
20 mM Tris-HCl (pH 8.0), 200 mM NaCl, 2 mM ATP, and 1 mM dithiothreitol(DTT) in the presence of [␥-32P]ATP (NEN) at 27°C for 1 h. As an enzymatic
source, a cytoplasmic lysate of S2 cells after centrifugation at 100,000 ⫻ g for 30
min was used for the experiments for which results are shown in Fig. 1B and 2.
Purified mammalian CKII (Fig. 6D) was a kind gift from U. Kikkawa (22). Afterthe reaction, proteins were separated on SDS-polyacrylamide gels and autoradiography was carried out to visualize phosphorylated proteins.
Mass spectrometry for identification of in vivo phosphorylation sites. Coomassie-stained dFMR1 bands were digested with trypsin (37), and the digest
solution was desalted with a ZipTip C18 tip (Millipore). The peptide mixture
eluted with 70% acetonitrile and 1% formic acid was analyzed by using a Q-Tof2
mass spectrometer (Micromass, Manchester, United Kingdom) equipped with ananoelectrospray ionization source, and the phosphopeptide sequence was obtained by tandem mass spectrometry (MS/MS). Mass spectra were collected inpositive-ion mode. Phosphopeptides were identified by MS data searching
against the National Center for Biotechnology Information (NCBI) database byusing the Mascot Search program (Matrix Science, London, United Kingdom).
MS/MS data were analyzed by the PepSeq program in the BioLynx software
package (Micromass) for determination of peptide sequences.
Purification and identification of dCKII. A cytoplasmic lysate of S2 cells
prepared as described above was resolved on a linear sucrose gradient (5 to30%). The gradient was centrifuged at 4°C in a Beckman MLA-130 rotor at
40,000 rpm for 90 min. Following centrifugation, fractions were collected andassayed for kinase activity toward GST-dFC285. The active fractions from thegradient were pooled, applied to a gel filtration PC-10 column (Amersham
Bioscience) to eliminate sucrose, and eluted with buffer A containing 10 mM
Tris-HCl (pH 7.5) and 100 mM NaCl prior to being applied to a UnoQ ionexchange column (Bio-Rad). The column was washed extensively with buffer A,
and a 100 mM to 1 M NaCl gradient of buffer A was applied to the column.
Fractions (0.5 ml) were collected and assayed for kinase activity. The phosphorylation-active fractions were pooled and diluted with 10 mM Tris-HCl (pH 7.5)
to a final 100 mM NaCl concentration prior to being applied to a UnoS ionexchange column (Bio-Rad). The column was washed extensively with buffer A,
and a 100 mM to 1 M NaCl gradient of buffer A was applied to the column.
Fractions (0.5 ml) were again collected and assayed for activity. The active
fractions were pooled and diluted with 10 mM Tris-HCl (pH 7.5) to a final 100
mM NaCl concentration prior to being applied to a heparin column. The column
was washed extensively with buffer A, and proteins were eluted stepwise with 10
mM Tris-HCl (pH 7.5) containing NaCl at the concentrations indicated in Fig.
4C. Each fraction was assayed for kinase activity and for protein content by silver
staining after being resolved on an SDS-polyacrylamide gel. Both protein bands(as indicated in Fig. 4C, lane 1 M NaCl) were sent to the Genomic ResearchCenter, Shimadzu Biotech, and mass spectrometric analysis was carried out toidentify them.
Production of recombinant alpha and beta subunits of dCKII. In order toobtain the cDNAs encoding the alpha and beta subunits of dCKII, poly(A)⫹RNA was purified from S2 cells and reverse transcription-PCR (RT-PCR) was
carried out using primers specifically designed for each clone (35). The cDNAsof the alpha and beta subunits of dCKII were subcloned into pGEX-5X andpET28 (Novagen) to produce GST-dCKII alpha and His-dCKII beta, respectively. Both fusion proteins were produced and purified as described by the
manufacturers.
Phosphorylation of hFMR1 in vivo. HeLa cells were grown as described
previously (42). HeLa cells transiently expressing myc-tagged hFMR1 were labeled with [32P]orthophosphate (1 mCi/ml) (Amersham Bioscience) in phosphate-free medium supplemented with 10% fetal calf serum. After incubationfor 3 h, the cells were lysed in phosphate-buffered saline buffer containing 1%
Empigen, 1 mM EDTA, 0.1 mM DTT, 2 ␮g of leupeptin/ml, 2 ␮g of pepstatin/
ml, and 0.5% aprotinin. After brief sonication, the lysate was clarified by centrifugation at 16,000 ⫻ g for 5 min and the supernatant was immunoprecipitatedwith either an anti-myc antibody (9E10) or a nonimmune control antibody(SP2/0). After extensive washing, the immunoprecipitates were separated on an
SDS-polyacrylamide gel and proteins labeled with 32P were visualized by using
BAS-2500 (FUJIFILM).

MATERIALS AND METHODS
Cell culture. S2 cells were grown in Schneider’s Drosophila medium (Gibco
BRL) supplemented with 10% fetal bovine serum (Sigma), 1% L-glutamine 50 Uof penicillin/ml, and 50 ␮g of streptomycin/ml (Gibco BRL) at 26°C. HeLa cells
were grown in Dulbecco’s modified Eagle medium (Sigma) supplemented with
10% fetal bovine serum, 1% L-glutamine, 50 U of penicillin/ml, and 50 ␮g ofstreptomycin/ml (Gibco BRL) at 37°C.
Subcellular fractionation, CIP treatment, and Western blot analysis. A cytoplasmic lysate of S2 cells was prepared in a lysis buffer containing 10 mM
Tris-HCl (pH 7.5), 150 mM NaCl, 50 ␮g of digitonin/ml, 2 ␮g of leupeptin/ml, 2
␮g of pepstatin/ml, and 0.5% aprotinin. To a half-portion of the lysate, which was
not to be treated with calf intestine alkaline phosphatase (CIP), NaF was added
to 2% immediately after preparation of the lysate. A cytoplasmic lysate from fly
ovaries was prepared in a similar lysis buffer, but 50 ␮g of digitonin/ml was
replaced by 0.5% Triton X-100. Treatment of the cytoplasmic lysates with CIP(NEB) was carried out at 30°C for 20 min. The lysates with and without CIP
treatment were then mixed with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer, and proteins were separated on an
SDS–7.5% polyacrylamide gel after heat denaturation. Western blotting was
carried out essentially as described previously (41) by using an anti-dFMR1
monoclonal antibody (48). Protein bands were visualized with an ECL Western
blotting detection kit (Amersham Bioscience).

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Protein-protein interaction assays. A fusion construct encoding dFMR1 and atandem affinity purification (TAP) tag (33) under the control of the Drosophila
metallothionein promoter in the pRmHa-3 vector (25) (a kind gift of F. Lafont)
was transfected into S2 cells. Expression of the dFMR1-TAP fusion protein was
induced by adding copper ions to the medium, and the fusion protein andassociated components were recovered from the cytoplasmic lysates of the S2cells by affinity selection on an immunoglobulin G (IgG) Sepharose 6 Fast Flowcolumn (Amersham Bioscience). After extensive washing with the binding buffer(20 mM Tris-HCl [pH 7.5], 0.5% Triton X-100, 150 mM NaCl, 2 ␮g of leupeptin/
ml, 2 ␮g of pepstatin/ml, and 0.5% aprotinin), the bound materials were separated on an SDS-acrylamide gel and analyzed by Western blotting using ananti-dFMR1 antibody. For the GST pulldown assay, GST-dCKII alpha, GSTdFMR1, and GST alone were bound to 30 ␮l of glutathione-Sepharose 4B resin(Amersham Bioscience) in 1 ml of binding buffer (20 mM Tris-HCl [pH 7.5],
0.5% Triton X-100, and 150 mM NaCl). Following incubation at 4°C for 1 h, theresins were pelleted and washed with the binding buffer. Either His-dCKII beta
or His-dFMR1 was then added to the resins and bound in the absence or in thepresence of MgCl2 at 10 mM and ATP at 1 mM at 4°C for 2 h. After extensive
washing with the binding buffer, the bound proteins were analyzed by Western
blotting to visualize His-dCKII beta and His-dFMR1 by using an anti-His antibody (Santa Cruz). A cytoplasmic lysate of S2 cells prepared as described above
was used for the in vivo interaction of dCKII with dFMR1. The antibodies
against rat CKII cross-reacting to dCKII (a kind gift from U. Kikkawa) werebound on GammaBind G Sepharose resin (Amersham Bioscience) and incubated with the S2 cytoplasmic lysate at 4°C for 1 h. After extensive washing of theresins, the bound proteins were analyzed by Western blotting using an antidFMR1 antibody. For Fig. 7, in vitro-phosphorylated GST-dFMR1 and HisdFMR1 were obtained as described above with a minor modification, namely,that [␥-32P]ATP was replaced with additional 2 mM ATP. In addition, the 1 mM
DTT in the buffer was replaced with 0.1 mM tris(2-carboxyethylphosphine)hydrochloride for HIS-dFMR1 phosphorylation. The enzymatic source used in theexperiments was the recombinant dCKII shown in Fig. 5A. Phosphorylated
GST-dFMR1 was bound to 30 ␮l of glutathione-Sepharose 4B resin in 1 ml ofbinding buffer (20 mM Tris-HCl [pH 7.5], 0.5% Triton X-100, and NaCl at theconcentration indicated). Following incubation at 4°C for 1 h, the resin waspelleted and washed with the binding buffer. His-dFMR1, either in a phosphorylated state or in an unphosphorylated state, was then added to the resin with
1 ml of the binding buffer and incubated at 4°C for 2 h. After extensive washingwith the binding buffer, the bound proteins were detected on Western blots byusing an anti-His antibody.
RNA-binding assay. Binding of His-dFMR1 in phosphorylated and unphosphorylated states was carried out by using 30 ␮l of either AGPOLY(U) type 6(Amersham Bioscience) or poly(G) immobilized on polyacrylhydrazido-agarose(Sigma) in 1 ml of binding buffer (20 mM Tris-HCl [pH 7.5], 0.5% Triton X-100,
and NaCl [Fig. 7C] or KCl [Fig. 7D] at the concentrations indicated) at 4°C for
2 h. After incubation, the beads were washed with the binding buffer. Boundproteins were then eluted by addition of SDS-PAGE sample buffer to the beads
and heat denaturation. Western blotting was performed to visualize His-dFMR1
by using an anti-His antibody.

RESULTSdFMR1 protein is phosphorylated in vivo and in vitro. In thecourse of experiments to elucidate the function of dFMR1, we
observed that dFMR1 in S2 cells migrates as several bands
when probed with an anti-dFMR1 antibody. This suggested
that dFMR1 is posttranslationally modified in vivo and thatone such modification is likely phosphorylation. In order to test
this hypothesis, the cytoplasmic lysate from S2 cells was treatedwith CIP, an enzyme known to dephosphorylate both nucleic
acids and proteins, and Western blot analysis was performed.
In the lysate without CIP, dFMR1 was detected as a strong
doublet (⬃75 kDa) and a weak upper band (⬃80 kDa) on an
SDS–7.5% polyacrylamide gel (Fig. 1A, ⫺CIP lane in S2 panel). After CIP treatment, however, the doublet became a discrete single band, apparently comigrating with the lower bandof the doublet in the untreated lanes (Fig. 1A, ⫹CIP lane in S2
panel), demonstrating that dFMR1 is phosphorylated in vivo
and that the phosphorylation state of dFMR1 is normally het-

FIG. 1. dFMR1 is phosphorylated in vivo and in vitro. (A) dFMR1proteins in cytoplasmic lysates of S2 cells and of fly ovaries before andafter CIP treatment were visualized by Western blotting using ananti-dFMR1 antibody (50). Migration changes of dFMR1 proteins areobserved before and after CIP treatment on SDS–7.5% polyacrylamide gels, indicating that dFMR1 proteins both in cultured cells andin animals are phosphorylated in vivo. (B) After production of GSTdFMR1, an in vitro phosphorylation assay was carried out using thecytoplasmic lysate of S2 cells in the presence of [␥-32P]ATP, and thephosphorylated proteins were visualized by autoradiography. The fulllength GST fusion protein is clearly phosphorylated, whereas GST
alone and the truncated by-product of GST-dFMR1 (⬃65 kDa in the
CBB panel) (see the text) are not, suggesting that a protein with adFMR1-phosphorylating activity exists in the cytoplasm of S2 cells andthat the activity is sequence specific. It is noted that EDTA abolishes
the activity. The left panel shows the protein substrates stained with
Commassie brilliant blue (CBB), and the positions of molecular mass
markers are indicated on the left of the gel.

erogeneous in the cells. After CIP treatment, the weak upperband also migrated slightly faster, but did not disappear, indicating that this weak band also represents dFMR1 in a phosphorylated form but that its larger size suggests other modifications and/or alternative splicing for dFMR1. The same
phenomenon was observed with fly ovaries (Fig. 1A, rightpanel), demonstrating that phosphorylation of dFMR1 is not
restricted to cultured cells but also occurs in animals. We next
investigated if dFMR1 could be phosphorylated in vitro. Recombinant full-length dFMR1 tagged with GST was purified
from Escherichia coli and mixed with a cytoplasmic lysate prepared from S2 cells in the presence of Mg2⫹ and [␥-32P]ATP.
As expected, GST-dFMR1 was observed to be phosphorylated
under such conditions, whereas GST alone, employed as anegative control, was not (Fig. 1B). When EDTA was added to
the reaction mixture to remove Mg2⫹, the modification was notobserved. Interestingly, the truncated peptide of GST-dFMR1(⬃65 kDa; CBB panel in Fig. 1B), which likely lacks the C
terminus of the fusion protein, since it was positive by Western

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FIG. 2. Determination of the phosphorylation site in dFMR1 by systematic mutation analyses. (A) The four parts of dFMR1 (shown in the
diagram at the bottom) were fused individually to GST and assayed for in vitro phosphorylation. It may be observed that only GST-dFC285,
containing 285 amino acids from the end of dFMR1, becomes phosphorylated, indicating that the phosphorylated region includes the RGG box.
In the schematic drawings, the locations of the first and the second KH domains, the Leu-rich region, and the RGG box are indicated as KH1,
KH2, Leu, and RGG, respectively. (B) To narrow down the phosphorylation region of dFMR1, the dFC285 fragment was further delineated fromits N terminus (shown in the diagram at the bottom), fused to GST, and assayed. GST-dFC277 was phosphorylated efficiently, whereas
GST-dFC240 was not, indicating that the region covering amino acid residues 405 to 441 in dFMR1 contains the phosphorylation site. (C) In theregion identified in panel B, each of four Ser residues was mutagenized to Ala in the context of GST-dFC285, and mutants were assayed todetermine the phosphorylation site in dFMR1. The phosphorylation level of the S406A mutant was detectable but was markedly reduced comparedto those of the others, demonstrating that Ser406 is the major phosphorylation site in dFMR1. (D) The amino acid sequence of the region
containing the phosphorylation site, Ser406 (circled and shaded), and the RGG box of dFMR1 (48) is shown. Five RGG sequences found in the
RGG box are indicated by boxes.

blotting using an anti-GST antibody (data not shown), was not
phosphorylated under the same circumstances. These results
suggest that the phosphorylation is sequence-specific and thatthe phosphorylation site is likely located towards the C terminus of dFMR1.
The phosphorylation site in dFMR1 is Ser406 preceding the
RGG box. To determine the phosphorylation site in dFMR1,
we decided to perform systematic mutation analyses. First,
dFMR1 was divided into four individual fragments, each ofwhich was fused to GST. After these proteins were purified
from E. coli, an in vitro phosphorylation assay was carried out.
We found that only one substrate, termed GST-dFC285, whichcontains 285 amino acid residues at the C terminus of dFMR1(amino acids 397 to the end), was phosphorylated in this assay(Fig. 2A). Since GST-dFC182, which contains 182 residues at
the C terminus of dFMR1 (amino acids 500 to the end), was
not phosphorylated, it was determined that a peptide corresponding to amino acids 397 to 499 of dFMR1 (corresponding
to the N-terminal region of the dFC285 fragment) contains thephosphorylation site. We next deleted stepwise the dFC285
fragment from its N terminus, fused it to GST, and performed
an in vitro phosphorylation assay. GST-dFC277 was phosphorylated as well as GST-dFC285 (Fig. 2B). In contrast, the phosphorylation of GST-dFC240, which lacks 45 residues at the N
terminus of dFC285, was markedly reduced (Fig. 2B), demonstrating that a peptide corresponding to amino acids 405 to 441
in dFMR1 contains the phosphorylation site and that a minorsite likely exists toward the end of the protein. We then alteredeach of four Ser residues within the region corresponding toamino acids 405 to 441 to Ala in the context of GST-dFC285.
Three point mutants (termed the S405A, S410A, and S413A
mutants) were phosphorylated to extents quite similar to thatof the wild type(Fig. 2C). Phosphorylation of S406A, on theother hand, was detectable, but the level was markedly reduced
compared to the others. Taken together, we determined that
Ser406 is the major phosphorylation site in dFMR1.
Ser406 of dFMR1 is phosphorylated in vivo. We used MS to
identify in vivo phosphorylation sites in dFMR1. dFMR1 was
immunoprecipitated from a cytoplasmic lysate prepared from
S2 cells, immunoprecipitates were resolved on an SDS-PAGE
gel, and dFMR1 was analyzed by MS (37). Trypsin digestionof the dFMR1 band identified a phosphopeptide spanning
Gly403 to Arg412 with a single phosphate group (data notshown), and MS/MS analysis of the phosphopeptide was used
to identify the exact phosphorylation site. The MS/MS spectrum of the phosphopeptide revealed the presence of a phosphate group at Ser406 (Fig. 3).
The protein phosphorylating dFMR1 is identical to dCKII.
To purify a protein with dFMR1-phosphorylating activity, acytoplasmic lysate of S2 cells was first fractionated on a linear
density sucrose gradient and assayed for phosphorylation activity toward GST-dFC285 (Fig. 4A). Fractions showing thehighest activity (F3, F4, and F5 in Fig. 4A) were pooled, andfurther purification was performed using various chromatog-

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FIG. 3. Ser406 of dFMR1 is phosphorylated in vivo. dFMR1 was immunoprecipitated from cytoplasmic lysates of S2 cells and resolved on an
SDS–7.5% PAGE gel. The band of dFMR1 was cut out and subjected to analysis of the phosphorylation site. Identification of phosphopeptides
was carried out using MS as described in Materials and Methods. The MS/MS spectrum of the tryptic phosphopeptide amino acids 403 to 412 ofdFMR1 obtained by collision-induced dissociation of the [M ⫹ 2H]2⫹ precursor ion, m/z 596.84, is shown. Product ions in the spectrum represent
mainly single-event preferential cleavage of the peptide bonds, resulting in simultaneous recording of sequence information from the N and C
termini (b- and y-type ions, respectively) of the peptide. This spectrum was analyzed by the PepSeq program in the BioLynx software package andwas matched to a dFMR1 peptide with additional mass from a phosphate residue (sequence shown on the top). With three potential sites ofphosphorylation (three serines), the correct assignment (Ser406) was unambiguously determined based on the presence of ions derived by cleavage
at the serine-serine peptide bond. This results in a y7 (m/z 885.43, 805.46 [peptide] ⫹ 79.97 [phosphate]) and a ⌬y7 (m/z 787.41) due to a neutral
loss (H2O ⫹ phosphate, 97.98) by ␤-elimination. ⌬y8 (m/z 874.43), due to a neutral loss (H2O ⫹ phosphate) of y8, was detected, and its phosphateshould be from Ser406.

raphy columns (Fig. 4B). Individual fractions eluted from thelast column (a heparin column) were assayed and separated on
an SDS-polyacrylamide gel, followed by silver staining to visualize the protein pattern (Fig. 4C). The fraction containing
kinase activity eluted at 1 M NaCl and contained two predominant proteins with apparent molecular masses of ⬃35 and ⬃28
kDa (Fig. 4C). Interestingly, these two proteins were observed
to exist in an approximately equal molar ratio, suggesting thatthey might comprise the enzyme that phosphorylates dFMR1.
The ⬃35- and ⬃28-kDa proteins in the fraction at 1 M NaCl
were isolated and subjected to MS analysis. We found that they
were identical to the alpha and beta subunits of dCKII (35),respectively (Fig. 4C).
Confirmation of dCKII as the protein phosphorylatingdFMR1 in vitro. To confirm that dCKII is indeed the protein
phosphorylating dFMR1, we obtained by RT-PCR the cDNA
clones encoding both the alpha and beta subunits of dCKII(35) and inserted them into vectors to produce the GST-dCKII
alpha and His-dCKII beta proteins (Fig. 5A). We first exam-

ined whether these two recombinant proteins can interact witheach other, as might be expected (see Discussion) (11, 35). We
found that GST-dCKII alpha interacted with His-dCKII beta
and that the affinity of the two proteins was stronger in thepresence of Mg2⫹ and ATP (Fig. 5B). Interaction between
dFMR1 and each subunit of dCKII was also examined. Eachsubunit interacted with dFMR1 independently of the other,and the affinity of the alpha subunit for dFMR1 was again
increased by addition of Mg2⫹ and ATP to the buffer (Fig. 5B).
We also examined if dCKII interacts with dFMR1 in vivo.
Immunoprecipitation from an S2 cytoplasmic lysate was first
performed with anti-rat CKII (rCKII) antibodies cross-reacting with dCKII (data not shown), and then Western blotting
was performed using an anti-dFMR1 antibody. A band corresponding to dFMR1 was observed in the lane with anti-rCKII
antibodies, but not with the nonimmune control antibody (SP2/0), demonstrating that dFMR1 indeed interacts with dCKII invivo (Fig. 5C). We next examined the phosphorylation activityof recombinant dCKII toward GST-dFC285. As expected, theVOL. 22, 2002

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hFMR1 is phosphorylated in cultured cells, in vivo 32P labelingof HeLa cells expressing myc-tagged hFMR1 was performed,
and myc-tagged hFMR1 was immunoprecipitated with antimyc antibodies. Autoradiography of the immunoprecipitates
separated by SDS-PAGE revealed that predominantly one
protein band was labeled with 32P and that it migrated at the
expected size of myc-tagged hFMR1 (⬃80 kDa) (Fig. 6B); thissuggests that hFMR1 is a phosphoprotein in vivo. To see if theresidue (Ser500) corresponding to Ser406 of dFMR1 (Fig. 6A)
is the phosphorylation site in hFMR1, the residue was altered
to Ala in the context of full-length hFMR1 and an in vitro
phosphorylation assay was carried out using recombinant
dCKII. As shown in Fig. 6C, a GST-hFMR1 mutant (S500A)
was not phosphorylated, whereas wild-type hFMR1 was.

FIG. 4. Purification and identification of dCKII as the protein
phosphorylating dFMR1. (A) The S2 cytoplasmic lysate with adFMR1-phosphorylating activity was applied to a linear sucrose density gradient (5 to 30%). After centrifugation, collected fractions wereassayed for phosphorylation toward GST-dFC285. Relatively high activities were observed in fractions F3 to F5, which were pooled andemployed for further purification steps (see the text). (B) Scheme forpurification steps after the sucrose gradient. The NaCl concentrations
at which the active fractions were eluted from each column are individually indicated. (C) As the final purification step, a heparin column
was used. After stepwise elution with the NaCl concentrations indicated, each fraction was assayed for phosphorylation and for protein
contents by silver staining after separation of the proteins on an SDSpolyacrylamide gel. The activity was concentrated in the fraction
eluted at 1 M NaCl, in which two proteins (⬃35 and ⬃28 kDa) wereprominent. Both were subjected to MS analysis and were found to be
identical to the dCKII alpha and beta subunits (35), respectively.

alpha subunit efficiently phosphorylated the substrate (see Discussion), whereas the beta subunit did not (Fig. 5D). When
GST-dFMR1 was substituted for GST-dFC285, the same data
were obtained (data not shown). Interestingly, addition of thebeta subunit to the reaction mixture together with the alphasubunit did not appear to alter the kinase activity of the alphasubunit (Fig. 5D). The contribution of the beta subunit todFMR1 phosphorylation remains unknown.
The Ser406 phosphorylation site in dFMR1 is highly conserved among FMR1/FXR family members across species. Determining whether FMR1/FXR family members from otherspecies are also phosphorylated by CKII was of great interest.
First, we inspected the amino acid sequences of those proteins(40, 47, 48, 52) and found that the region surrounding Ser406
in dFMR1 is highly conserved (Fig. 6A). To analyze whether

FIG. 5. Confirmation of dCKII as the protein with dFMR1-phosphorylating activity. (A) After the cDNAs of dCKII alpha and beta
subunits were obtained by RT-PCR, they were subcloned into bacterial
expression vectors, and GST-dCKII alpha and His-dCKII beta were
produced. Both purified proteins were analyzed by staining with Coomassie brilliant blue on an SDS-polyacrylamide gel. The positions ofmolecular mass markers are indicated on the left of the gel. (B) Interaction between the dCKII subunits and dFMR1 in vitro. Specificinteraction is observed between GST-dCKII alpha and His-dCKII beta
or His-dFMR1. In both cases the stronger protein-protein interactions
are seen in the presence of MgCl2 and ATP. Not only the alpha but
also the beta subunit interacts independently of the other with GSTdFMR1. (C) In vivo interaction of dCKII with dFMR1. Immunoprecipitation was performed from an S2 cytoplasmic lysate using antirCKII antibodies and a nonimmune control antibody (SP2/0). Western
blotting of the immunoprecipitates using an anti-dFMR1 antibody
revealed that dFMR1 specifically coimmunoprecipitated with dCKII.
(D) The phosphorylation activities of the recombinant proteins toward
GST-dFC285 were assayed. GST-dCKII alpha (shown in panel A)
efficiently phosphorylates the substrate, whereas His-dCKII beta has
no such activity. It should be noted that the beta subunit added together with the alpha subunit in the reaction mixture did not influence
the activity of the alpha subunit itself.

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MOL. CELL. BIOL.

associated components were recovered from cell extracts byaffinity selection on an IgG matrix and were then analyzed by
SDS-PAGE and Western blotting. As shown in Fig. 7A, notonly the TAP-tagged dFMR1 but also endogenous dFMR1

FIG. 6. Phosphorylation of FMR1/FXR in humans. (A) The peptide sequence containing S406 of dFMR1 is aligned with the corresponding regions in hFMR1 (47), Xenopus FMR1 (xFMR1) (39), zebra fish FMR1 (zFMR1) (48), human FXR1 (hFXR1) (39), Xenopus
FXR1 (xFXR1) (39), and human FXR2 (hFXR2) (52). The phosphorylated residue (S406) in dFMR1 is highly conserved among the sequences and is highlighted on a solid background. Other highly conserved residues are shaded. (B) Phosphorylation of hFMR1 in vivo.
HeLa cells 24 h after transfection of myc-hFMR1 were labeled with
32
P. Immunoprecipitations were performed with an anti-myc monoclonal antibody (9E10) and a nonimmune control antibody (SP2/0). Autoradiography of the immunoprecipitates separated by SDS-PAGE
revealed that one protein band was predominantly labeled with 32P andmigrated at the expected size of myc-tagged hFMR1 (⬃80 kDa). Thisband was observed only in the 9E10 immunoprecipitates. (C) The
predicted phosphorylation site in hFMR1 (Ser500) was mutagenized
to Ala in the context of full-length GST-hFMR1, and an in vitro
phosphorylation assay was carried out using the recombinant dCKII
shown in Fig. 5A and mammalian CKII (indicated as rCKII). The
GST-hFMR1 mutant (S500A) is not phosphorylated, in contrast towild-type (wt) GST-hFMR1, demonstrating that the conserved Ser
residue is the phosphorylation site in hFMR1.

Ser500 in hFMR1 was, therefore, determined to be the phosphorylation site of hFMR1, and there seem to be no otherphosphorylation sites in this case. We also tried purified rCKII
in the assay and found that the mammalian enzyme phosphorylated wild-type hFMR1, but not the S500A mutant (Fig. 6C).
Moreover, we also observed that the mammalian CKII phosphorylated dFMR1 (data not shown).
Phosphorylation of dFMR1 modulates the efficiency of
forming homomers and of binding RNA in vitro. It has beenreported that dFMR1 forms homomers in vitro (48), just as
mammalian FMR1/FXR proteins do (52). To address thequestion of whether dFMR1 also forms homomers in vivo, weused the TAP tag method (33). The TAP-tagged dFMR1 protein was expressed in S2 cells, and the fusion protein and

FIG. 7. Biochemical properties of dFMR1 are altered by dCKII
phosphorylation. (A) An in vivo protein-protein interaction assay was
carried out using the TAP tag method. A fusion construct encoding thecalmodulin-binding peptide, a TEV protease recognition sequence,
two IgG-binding units of protein A of Staphylococcus aureus, anddFMR1 under the control of the Drosophila metallothionein promoter
was transfected into S2 cells. The fusion protein and associated components were recovered from cell extracts by affinity selection on an
IgG matrix. After extensive washing, the bound material was analyzed
on a gel, and Western blotting using anti-dFMR1 antibodies was performed. (B) An in vitro protein-protein interaction assay was carriedout. Phosphorylated and unphosphorylated His-dFMR1s (input) wereincubated with GST-dFMR1 in a phosphorylated state immobilized on
glutathione-Sepharose 4B resin at the NaCl concentrations indicated.
After extensive washing, the bound fraction was analyzed by Western
blotting using an anti-His antibody. (C) An in vitro RNA-binding
assay was carried out using poly(U) beads. Phosphorylated andunphosphorylated His-dFMR1s (input) were incubated withpoly(U) at the NaCl concentrations indicated. After extensive washing, the bound fraction was analyzed by Western blotting as forpanel B. (D) An in vitro RNA-binding assay was carried out using
poly(G) beads. In this experiment, KCl was used instead of NaCl. The
unbound fractions are shown below in comparison with the input
diluted 1:10, which clearly shows the ratio of the unphosphorylated tothe phosphorylated proteins. (E) Poly(G)-binding comparison in thepresence of K⫹ and Li⫹. Phosphorylated and unphosphorylated HisdFMR1s were incubated with poly(G) in the presence of NaCl and
LiCl, both at 150 mM. After extensive washing, the bound fraction was
analyzed by Western blotting as for panel D.

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PHOSPHORYLATION OF FMR1 PROTEIN BY CKII

was found in the IgG-bound material, indicating that dFMR1forms homomers in vivo. We then tried to ascertain if thephosphorylation affects the oligomerization of dFMR1. For
this experiment, GST- and His-tagged dFMR1s purified from
E. coli were phosphorylated by recombinant dCKII. That mostof the population of GST- and His-dFMR1 proteins were
phosphorylated was confirmed by observing the mobilitychange on Western blots (see Fig. 7B, input lanes, for HisdFMR1). GST-dFMR1 in a phosphorylated state (data notshown) was first bound to glutathione-Sepharose resins, andafter extensive washing, a GST pulldown assay was carried out
at 150 and 450 mM NaCl. By performing Western blotting
using an anti-His antibody, it was clearly observed that thephosphorylated His-dFMR1 bound much more efficiently thanthe unphosphorylated His-dFMR1 (Fig. 7B), demonstratingthat the oligomerization of dFMR1 is upregulated by dCKII
phosphorylation. We next investigated whether the RNA-binding activity of dFMR1 was also influenced by dCKII modification. When the assay was carried out with poly(U) beads,
phosphorylated dFMR1 showed a higher affinity for them than
did unphosphorylated dFMR1 (Fig. 7C). Considering the result shown in Fig. 7B, the phosphorylated dFMR1 likely forms
homomers more efficiently than the unphosphorylated dFMR1
at the NaCl concentrations used in this experiment. Thus, it isspeculated that phosphorylation followed by oligomerization
confers a higher RNA-binding activity on dFMR1 protein. We
also examined dFMR1 binding to poly(G) beads. Recently, it
was reported that dFMR1 preferentially binds G quartets in
RNA (7, 36). The G quartet is a secondary structure formed in
G-rich sequences and is known to be stabilized by certain
cations such as K⫹. Poly(G) is known to form G quartets (23).
Thus, this particular experiment was performed in a buffer
containing KCl. In this assay, phosphorylated and unphosphorylated dFMR1s showed approximately the same affinity forpoly(G) at any given concentration of KCl (Fig. 7D). Interestingly, it was clearly observed that the unphosphorylated formof dFMR1 coexisting with the phosphorylated form in the
input (see the right input lane in Fig. 7D) was not detected in
the bound fraction but was observed in the unbound fraction(Fig. 7D). When the unphosphorylated form was allowed tobind to the beads first and then the phosphorylated form wasadded, both forms were detected in the bound fraction (datanot shown), suggesting that phosphorylated dFMR1 does nothave the ability to displace unphosphorylated dFMR1 frompoly(G) beads. It is known that G quartets are stabilized moreby K⫹ than by Li⫹ and that hFMR1 shows a higher affinity fora G quartet formed in the presence of K⫹ (36). Thus, thepoly(G)-binding activities of phosphorylated and unphosphorylated dFMR1s were compared in the presence of K⫹ and Li⫹.
Interestingly, the unphosphorylated form of dFMR1 showedno significant difference between K⫹ and Li⫹, whereas thephosphorylated form showed much higher affinity for poly(G)in a buffer containing K⫹ (Fig. 7D), suggesting that in binding
to a G quartet, the phosphorylated dFMR1 behaves as does
hFMR1.

that bacterially produced GST-dFMR1 is phosphorylated witha cytoplasmic lysate from S2 cells in vitro. We have also shownthat the protein with dFMR1-phosphorylating activity purified
from the lysate was dCKII (35) and that recombinant dCKII
does indeed phosphorylate dFMR1 in vitro.
CKII is known to phosphorylate both nuclear and nonnuclear proteins (46, 50) and is highly conserved from yeasts tohumans. The high degree of conservation suggests that CKII
has an essential role in organisms. Indeed, genetic analyses inyeast have demonstrated that the enzyme is essential for viability (32, 44). We found in this study that at least in vitro, bothdCKII and mammalian CKII phosphorylate both dFMR1 andhFMR1 at the sites we identified by performing mutation analyses. The cross-species activity implies that the phosphorylation of FMR1 proteins is highly conserved through evolution
and is important, perhaps even required, for function of the
FMR1/FXR proteins in divergent organisms.
CKII is composed of alpha and beta subunits that are associated as a heterotetramer (␣2␤2) (22). The alpha subunit
contains a protein kinase domain that is homologous to thecatalytic domain of other protein kinases (15). The beta subunit has been proposed to play a complex role in regulating thebasal activity of the alpha subunit (4). Recently, the physicalinteraction of the CKII enzyme and its substrates was shown to
be mediated by the beta, but not the alpha, subunit (10). In thepresent study, we observed that both the beta and the alphasubunits are able to interact directly with dFMR1 in vitro andthat the affinity of the alpha subunit for dFMR1 is increased in
the presence of Mg2⫹ and ATP. We also found that the alphasubunit is necessary and sufficient for efficient phosphorylationof dFMR1 in vitro; that is, the beta subunit seemed not to
influence that activity. The role of the beta subunit of CKII in
the phosphorylation of FMR1 remains undefined.
CKII phosphorylates Ser/Thr residues of many protein substrates in vivo, and the consensus sequence is known to be(Ser/Thr)-(Asp/Glu)-X-(Asp/Glu) (24). We determined thatthe peptide sequence in dFMR1 containing the phosphorylation site (Ser406) was Ser-Asp-Ile-Glu, which perfectly fits theconsensus. The phosphorylation site is highly conserved within
FMR1/FXR family members from flies to humans. The sequences of these other family members also match the CKII
consensus. We have mentioned that there might be minor
phosphorylation sites in dFMR1, since the S406A mutation did
not result in a total loss of phosphorylation. However, there
are no other sequences in dFMR1 that perfectly fit the consensus sequence for CKII phosphorylation. In the case ofhFMR1, we could find two other sites that fit the consensus(namely, Thr455 and Thr518). But neither of these is likely to
be a site of modification by CKII, at least in vitro, since the
GST-hFMR1 mutant (S500A) that contains intact Thr455 andThr518 residues is not phosphorylated by either dCKII or
mammalian CKII. It remains possible, however, that other,
unidentified kinases or even CKII itself contributes phosphorylation of the other sites of hFMR1 in vivo. It is known that thenegative-charge requirement within the CKII consensus can besupplied by phosphorylated residues (46). If so, once Ser500 in
hFMR1 is phosphorylated, Ser497 and consequently Ser494
become appropriate targets for CKII. This sequential phosphorylation may have occurred even in vitro (see Fig. 6C), but
at this time we do not have data to support this scenario.

DISCUSSION
In this study we have shown that dFMR1 protein is phosphorylated in vivo, both in cultured cells and in animals, and

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Although it is understood that the phosphorylation activityof CKII appears to be constitutive, there are several studies
available showing the regulation of CKII phosphorylation activity in vivo. Recently, it was demonstrated that CKII phosphorylates the human p53 protein in response to UV irradiation and that the chromatin transcriptional elongation factor,
which forms a complex with CKII, alters the specificity of CKII
to selectively phosphorylate p53 over other substrates including casein (20). In the case of Dishevelled (Dsh), anotherexample of a substrate for dCKII, phosphorylation is regulated
in response to Dsh and Dfz2 (a Wingless receptor) expression
in the signal transduction pathway of the Wingless protein(50). When we used a linear density sucrose gradient as a firstmeans to purify dCKII, a relatively high activity was obtainedin fractions F3 to F5 (see Fig. 4A). This indicated that it was
likely that dCKII associates with other molecules, constituting
a relatively heavy complex. However, the complex may not bevery stable and probably is dissociated through a series ofpurification procedures, since no other proteins eluted withdCKII from the last heparin column (see Fig. 4C). At thispoint, we have no data identifying molecules that associatewith dCKII. It is possible, as in the case of p53, for example,
that the dCKII-interacting proteins and/or nucleic acids regulate the dCKII activity in the phosphorylation of dFMR1. We
are currently attempting to isolate the dCKII interactors andunderstand their involvement in the phosphorylation ofdFMR1.
It has been reported that the DNA-binding activity of somenuclear proteins is regulated by CKII phosphorylation (3, 28,
29, 45). In this study we have shown that the RNA-binding
activity of dFMR1 to poly(U) is upregulated by phosphorylation by dCKII in vitro. This property may be a result of thefavorable homotypic interactions of dFMR1 caused by phosphorylation (see Fig. 7B). We speculate that phosphorylationof dFMR1 induces a conformational change that results in astronger association with itself and perhaps also with poly(U).
Stable oligomerization of dFMR1 simply means that the KH
domains and the RGG boxes are also multiplied and arebrought close enough together to cooperate, which we think
confers the stronger ability to bind poly(U) beads. Surprisingly,
different results were obtained when we used poly(G) beads. In
such experiments, dFMR1, whether phosphorylated or unphosphorylated, showed approximately the same ability to bindpoly(G). On the other hand, when both proteins were added tothe beads together, only phosphorylated dFMR1 was detectedin the bound fraction. In different experiments, however,where unphosphorylated dFMR1 was allowed to bind poly(G)first, and phosphorylated dFMR1 was added later, both proteins were observed in the bound fraction. One explanation forthese observations is that the on-rate and off-rate of phosphorylated or unphosphorylated dFMR1 for poly(G) binding may
differ from each other. Data to support this speculation are
currently not available, and further detailed studies will benecessary.
These studies constitute the first demonstration that FMR1
is phosphorylated in vivo and that the phosphorylation alters
its biochemical properties, such as its interactions with itself
and with RNA in vitro. Although the in vivo relevance of theseobservations is currently unknown, our results should be the

MOL. CELL. BIOL.

starting point for a better understanding of the effects of phosphorylation on the function of FMR1.
ACKNOWLEDGMENTS
We thank Gideon Dreyfuss and Lili Wan for generously providing
dFMR1 cDNA and the anti-dFMR1 antibody, Ushio Kikkawa for
purified mammalian (rat) CKII and the anti-CKII antibody, Bertrand
Seraphin for the TAP plasmids, and Frank Lafont for pRmHa-3. We
gratefully acknowledge Eiji Majima for help with peptide sequencing
by nanoelectrospray MS. We also thank Paul Eder and Tetsuya Taurafor critical reading of the manuscript. We are grateful to members of
our laboratory, especially Hitomi Ichinose and Yoshinori Kawamura,for technical assistance.
This work was supported by grants from the FRAXA Research
Foundation, the Cure Autism Now (CAN) Foundation, the Japan
Society for the Promotion of Science (JSPS), the Ministry of Health,Labour and Welfare of Japan, and the Ministry of Education, Science,Sports, and Culture of Japan.
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