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MOLECULAR AND CELLULAR BIOLOGY, July 2010, p. 3286–3298 Vol. 30, No. 13
0270-7306/10/$12.00 doi:10.1128/MCB.01520-09
Copyright © 2010, American Society for Microbiology. All Rights Reserved.
PHF8 Targets Histone Methylation and RNA Polymerase
II To Activate Transcription
䌤
†
Klaus Fortschegger,
1
§‡ Petra de Graaf,
2
§ Nikolay S. Outchkourov,
2
§ Frederik M. A. van Schaik,
2
H. T. Marc Timmers,
2
* and Ramin Shiekhattar
1,3
*
Center for Genomic Regulation, Barcelona Biomedical Research Park, 08003 Barcelona, Spain
1
; Department of Physiological Chemistry,
University Medical Centre, 3584 CG Utrecht, Netherlands
2
; and Wistar Institute, Philadelphia, Pennsylvania 19104
3
Received 24 November 2009/Returned for modification 30 December 2009/Accepted 14 April 2010
Mutations in PHF8 are associated with X-linked mental retardation and cleft lip/cleft palate. PHF8 contains
a plant homeodomain (PHD) in its N terminus and is a member of a family of JmjC domain-containing
proteins. While PHDs can act as methyl lysine recognition motifs, JmjC domains can catalyze lysine de-
methylation. Here, we show that PHF8 is a histone demethylase that removes repressive histone H3 dimethyl
lysine 9 marks. Our biochemical analysis revealed specific association of the PHF8 PHD with histone H3
trimethylated at lysine 4 (H3K4me3). Chromatin immunoprecipitation followed by high-throughput sequenc-
ing indicated that PHF8 is enriched at the transcription start sites of many active or poised genes, mirroring
the presence of RNA polymerase II (RNAPII) and of H3K4me3-bearing nucleosomes. We show that PHF8 can
act as a transcriptional coactivator and that its activation function largely depends on binding of the PHD to
H3K4me3. Furthermore, we present evidence for direct interaction of PHF8 with the C-terminal domain of
RNAPII. Importantly, a PHF8 disease mutant was defective in demethylation and in coactivation. This is the
first demonstration of a chromatin-modifying enzyme that is globally recruited to promoters through its
association with H3K4me3 and RNAPII.
Posttranslational modifications of histone tails play an im-
portant role in chromatin structure and function (27). While
the presence of several modifications correlates with gene ac-
tivation and transcription, others have opposing repressive
functions and are enriched in transcriptionally inactive or het-
erochromatic regions. A well-studied type of histone modifi-
cation is methylation of lysine residues, which is conferred by
histone methyltransferases (KMTs). There are three possible
states of lysine methylation, namely, mono-, di-, and trimethyl-
ation. As expected for reversible marks involved in dynamic
gene regulation, the methyl groups can be removed by histone
demethylases (KDMs) of either the LSD1 or the Jumonji C-
terminal-containing (JmjC) family of proteins (43, 48).
The JmjC family of histone demethylases consists of approx-
imately 30 members in humans (9, 25). Plant homeodomain
(PHD) finger-containing proteins 2 and 8 (PHF2 and PHF8,
respectively) and KIAA1718 constitute a subgroup containing
a single N-terminal PHD followed by the catalytic JmjC do-
main. Several protein domains (including the PHD; the MBT,
WD40, and Tudor domains; and chromodomains) have been
shown to bind to peptides containing either an unmodified or
a methylated lysine residue, and some PHDs specifically inter-
act with histone H3 trimethylated at lysine 4 (H3K4me3) (42,
51, 53). H3K4me3 is considered to be an activating chromatin
mark because it is enriched at active RNA polymerase II
(RNAPII) transcription start sites (TSSs) (4, 17). In mamma-
lian cells, H3K4me3 is mainly established by the SET1/KMT2
family, which includes the mixed-lineage leukemia 1 to 4
(MLL1 to -4) proteins. These chromatin modifiers can be re-
cruited to promoters upon gene activation (16, 52), and certain
MLL complexes also contain KDMs that concomitantly re-
move the repressive H3K27me3 mark and enhance expression
(7, 21, 30).
On the other hand, H3K4me3 can be erased by KDMs of the
JARID1/KDM5 family that are part of polycomb-repressive
complexes (PRC1/2) (29, 35). These complexes also contain
KMTs that are able to methylate histone H3 at lysine 9 or 27.
Methylated H3K9 and H3K27 are repressive marks, which are
read by PRC1 via chromodomain-containing proteins, a mech-
anism that is thought to be responsible for enforcement and
inheritance of silenced chromatin (5, 18). This obvious cou-
pling of writers (KMTs), readers (like proteins containing
PHDs), and erasers (KDMs) enables cross talk between dif-
ferent chromatin modifications (45). Furthermore, many SET
and JmjC domain proteins contain additional domains or pu-
tative DNA-binding modules, increasing their specificity and
affinity for modified chromatin.
PHF8 is a ubiquitously expressed nuclear protein whose
dysfunction is implicated in disease (28). Mutations in
the PHF8 locus on the X chromosome have been linked to
Siderius-Hamel syndrome, an X-linked mental retardation
(XLMR) that is often accompanied by cleft lip and/or cleft
palate (44). The described mutations result mostly in early
truncations of the protein before or in the JmjC domain (1,
* Corresponding author. Mailing address for Ramin Shiekhattar:
The Wistar Institute, 3601 Spruce Street, Philadelphia, PA 19104.
Phone: (215) 898-3896. Fax: (215) 898-3986. E-mail: shiekhattar
@wistar.org. Mailing address for H. T. Marc Timmers: Department of
Physiological Chemistry, University Medical Centre, Universiteitsweg
100, 3584 CG Utrecht, Netherlands. Phone: (31) 88 756 8981. Fax: (31)
88 756 8101. E-mail: H.T.M.Timmers@umcutrecht.nl.
‡ Present address: Children’s Cancer Research Institute, St. Anna
Kinderkrebsforschung, Zimmermannplatz 10, 1090 Vienna, Austria.
§ K.F., P.D.G., and N.S.O. contributed equally to this work.
† Supplemental material for this article can be found at http://mcb
.asm.org/.
䌤
Published ahead of print on 26 April 2010.
3286
28). Moreover, a single point mutation (F279S) in the JmjC
domain and a microdeletion of the whole locus were also
reported to cause the disease phenotype (26, 36).
In the present study, we delineate crucial biochemical prop-
erties of PHF8. We report that PHF8 is a demethylase specific
for H3K9me2 and a binder of H3K4me3-marked nucleosomes.
Genome localizations show that PHF8, H3K4me3, and RNA-
PII cooccupy thousands of promoters. Association studies sug-
gest that PHF8 interacts directly with the C-terminal domain
(CTD) of the RPB1 subunit of RNAPII. The F279S mutant of
PHF8 displays cellular mislocalization, defective histone de-
methylation, and aberrant coactivation function.
MATERIALS AND METHODS
Vectors. Full-length PHF8 (NCBI reference sequences NM_15107.1 for
mRNA and NP_055922.1 for protein) was cloned into the pFlag-CMV2 vector
for expression in mammalian cells and into pFastBacHTa-Flag for expression in
insect cells. Truncated versions (containing amino acids 1 to 352 or 1 to 489 of
PHF8) were cloned into pFlag-CMV2 with a short simian virus 40 (SV40)
nuclear localization signal (NLS) (PKKKRKVG) added to the C terminus to
ensure correct nuclear import. The mutations D28A/W29A, H31A/C34A,
H247A/D249A, and F279S were generated by site-directed mutagenesis using
the QuikChange protocol (Stratagene). The cloning primer and mutagenesis
oligonucleotide sequences are described in the supplemental material.
JARID1A- and JMJD3-HA-pCMV plasmids were kind gifts from Kristian He-
lin’s laboratory. A PCR fragment encoding the PHD of human PHF8 (residues
2 to 66) (PHF8-PHD) was inserted into the pGEX-2T-derived pRPN265NB
plasmid. Glutathione S-transferase (GST)–PHF8-PHD mutant plasmids were
obtained by site-directed mutagenesis using the QuikChange protocol (Strat-
agene). The plasmids for GST-Taf3 (amino acids 857 to 924) and GST-UbcH5B
were described previously (14, 51). The EB2 plasmid encoding GST fused to 27
repeats of the consensus CTD (YSPTSPS) was a kind gift of Marc Vigneron
(ESBS, Strasbourg, France). All inserts were verified by DNA sequencing.
Recombinant protein expression and purification. Full-length PHF8 contain-
ing N-terminal Flag and His
6
tags was expressed in Sf9 cells using baculovirus
(Invitrogen Bac-to-Bac system). The cells were lysed in whole-cell lysis buffer (20
mM Tris-HCl, pH 7.4, 137 mM NaCl, 1 mM EDTA, 1.5 mM MgCl
2
, 10%
glycerol, 1% Triton X-100, 0.2 mM phenylmethylsulfonyl fluoride [PMSF], 1
g/ml aprotinin, 1 g/ml leupeptin, 1 g/ml pepstatin) 2 days after infection and
incubated with M2 Flag agarose beads (Sigma). The beads were washed four
times with BC500 buffer (20 mM Tris-HCl, pH 7.4, 0.2 mM EDTA, 10 mM
-mercaptoethanol, 10% glycerol, 500 mM KCl, 0.1% NP-40, 0.2 mM PMSF, 1
g/ml aprotinin, 1 g/ml leupeptin, 1 g/ml pepstatin), and recombinant protein
was eluted with Flag peptide (400 g/ml; Sigma) in BC500. After concentration
of the eluates by centrifugation in Microcon YM-10 columns (Millipore), frac-
tions were run on 4 to 12% Tris-glycine SDS-polyacrylamide gels and Coomassie
stained (Invitrogen), and the protein concentration was estimated by comparison
to bovine serum albumin (BSA) standards.
In vitro demethylation assay. Four micrograms of bulk histones from calf
thymus (Sigma) was incubated in 30 l demethylation buffer [20 mM Tris-HCl,
pH 7.4, 2 mM ascorbic acid, 1 mM ␣-ketoglutarate, approximately 100 mM KCl,
100 mM NaCl, 50 M (NH
4
)
2
Fe(SO
4
)
2
] with or without up to 10 g recombinant
PHF8 for4hat37°C. After denaturation in SDS loading buffer, the reaction
mixtures were run on 4 to 20% Tris-glycine SDS-polyacrylamide gels and blotted
to polyvinylidene difluoride (PVDF) membranes (Millipore). Western blots were
blocked for ⬎1 h at room temperature with 5% skim milk powder diluted in
Tris-buffered saline containing 0.1% Tween 20 (TBS-T) before sequential incu-
bation for1hatroom temperature with antibodies. Primary mouse or rabbit
antihistone antibodies were usually diluted 1:1,000, and secondary anti-IgG–
alkaline phosphatase antibodies were diluted 1:10,000 (Promega) in TBS-T. Sig-
nals were developed by incubation with 5-bromo-4-chloro-3-indolylphosphate–
nitroblue tetrazolium (BCIP-NBT) substrate, and the blots were dried and
scanned. All antibodies used in this study are listed in the supplemental material.
Cell culture and indirect immunofluorescence. 293T, U2OS, Hs68, and
HeLa-S3 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM)
containing 10% fetal bovine serum (FBS), L-glutamine, and antibiotics. Tran-
sient transfection of plasmid DNA was performed 6 h after seeding using Fu-
Gene6 transfection reagent according to the manufacturer’s instructions
(Roche). For immunofluorescence assays, 293T cells were seeded onto glass
coverslips and transfected with 1 g PHF8(1-489)NLS-pFlag-CMV2 wild-type or
mutant constructs using 3 l Metafectene Pro transfection reagent (Biontex) per
coverslip. Twenty-four to 48 h later, the coverslips were washed with phosphate-
buffered saline (PBS) and fixed in fresh 1% formaldehyde-PBS for 10 min. After
being washed, the cells were permeabilized in 0.1% Triton X-100–PBS for ⬎10
min and sequentially incubated for1hatroom temperature with antibodies.
Primary mouse or rabbit antihistone antibodies were diluted 1:200 to 1:500, and
anti-Flag antibodies were diluted 1:1,000 in 0.1% Triton X-100–PBS. Secondary
anti-mouse IgG–Alexa Fluor-488 and anti-rabbit IgG–Alexa Fluor-568 antibod-
ies (Invitrogen) were diluted 1:1,000 in 0.1% Triton X-100–PBS. After DAPI
(4⬘,6-diamidino-2-phenylindole) counterstaining, the coverslips were washed
with PBS and mounted on slides using Mowiol 4-88 containing 2.5% 1,4-
diazabicyclo[2.2.2]octane (DABCO) as an antibleach. Detailed information on
image acquisition and statistical analysis is given in the supplemental material.
ChIP. 293T, HeLa-S3, or Hs68 cells were trypsinized from 2 (293T and HeLa)
or 16 (Hs68; 4-day serum starved or 4-h serum restimulated) subconfluent 15-cm
dishes, washed with PBS, and fixed in 10 ml of 1% formaldehyde-PBS at room
temperature for 10 min. One milliliter of 1.25 M glycine was added and incu-
bated for 5 min at room temperature. The cells were pelleted, washed with cold
PBS, and lysed for 15 min on ice in 300 l chromatin immunoprecipitation
(ChIP) lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris-HCl, pH 8, and
protease inhibitors). After the addition of 1.7 ml ChIP dilution buffer (0.01%
SDS, 1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl, pH 8, 167 mM NaCl,
and protease inhibitors), chromatin was sonicated on ice in a Bioruptor (Diage-
node) for 10 min (30 s on, 30 s off; high power load). After dilution with another
1 ml of ChIP dilution buffer, solubilized chromatin was cleared and precleared by
incubation with protein A-agarose beads (Millipore). Five hundred to 1,000 lof
this chromatin solution (25 to 50 g DNA) was incubated overnight with 2 to 4
g of antibody (listed in the supplemental material). Protein A-bead slurry (30
l) was added, rotated for2hat4°C, and washed 3 times with low-salt buffer
(0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl, pH 8, 150 mM
NaCl), one time with high-salt buffer (0.1% SDS, 1% Triton X-100, 2 mM
EDTA, 20 mM Tris-HCl, pH 8, 500 mM NaCl), one time with LiCl buffer (250
mM LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA, 10 mM Tris-HCl, pH 8),
and one time with TE buffer (10 mM Tris-HCl, pH 7.6, 1 mM EDTA). Bound
chromatin was eluted for ⬎1 h at room temperature in 100 l fresh elution buffer
(0.1 M NaHCO
3
, 1% SDS). Cross-links were reversed overnight at 65°C, and
DNA was purified using a QIAquick PCR purification kit (Qiagen).
For ChIP followed by high-throughput sequencing (ChIP-seq), approximately
10 ng immunoprecipitated DNA per sample was used for end cleanup and
adaptor ligation. Size-selected preamplified inserts of 135 ⫾25 bp were se-
quenced by the Illumina/Solexa technique. Total reads were mapped to the
human genome version 18 using Paolo Ribeca’s GEM mapper (http://www
.paoloribeca.net/software/GEM/index.html) (see Table S2 in the supplemental
material). Non-strand-specific cluster/peak analyses were performed with the
NGS-Analyzer (Genomatix RegionMiner 2.02) using a 100-bp window and a
Poisson distribution-derived cutoff value (depending on total reads; at least 8 to
10 reads per cluster). Regions overlapping with clusters of irrelevant IgG ChIP
(for HeLa and 293T cells) or input DNA control (for Hs68 cells) were filtered
out. Overlap analyses were done with Genomatix GenomeInspector (Eldorado
12-2008) using our own ChIP-seq data and published HeLa data for H3K4me3
(37) and for RNAPII (38). Strand-specific read analyses were performed with the
ChIP-cor tool (http://ccg.vital-it.ch/chipseq/). Reads were correlated to oriented
TSSs (49,392 ENSEMBL50 features). The examined distance from the reference
feature was ⫾1,000 bp, the window width was 20 bp, the cutoff value was 10
counts, and the output format was normalized counts. Wiggle files for display in
the UCSC genome browser were generated using the ChIP-center tool (hg18; tag
shift, 60 bp; count cutoff, 10; resolution, 20 bp). For correlation studies, pub-
lished HeLa expression data were used (10).
For quantitative ChIP (qChIP), purified immunoprecipitated DNA was used
as a template for real-time PCR with 0.2 M locus-specific primers (sequences
are given in the supplemental material) and Bio-Rad IQ SYBR green mix.
Samples and dilution series of input DNA were run in triplicate. The immuno-
precipitated percentage of input was calculated using individual standard curves.
Bacterial protein expression and pulldowns. Expression of GST fusion pro-
teins was induced, and lysates were prepared essentially as described previously
(14). H3 peptides (⬃0.1 nmol) were coupled to magnetic streptavidin beads
(M-280; Dynal) in H3-binding buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl,
0.1% NP-40, 10 M ZnCl
2
, 1 mM dithiothreitol [DTT], and protease inhibitors),
and excess peptide was removed by washing. Crude bacterial lysates were incu-
bated with peptide-coated streptavidin beads in binding buffer (50 mM Tris-HCl,
pH 8, 150 mM NaCl, 0.1% NP-40, 10 M ZnCl
2
, 1 mM DTT, and protease
inhibitors) for 2 to3hat4°C. Following extensive washing, bound proteins were
VOL. 30, 2010 PHF8 IS A TRANSCRIPTIONAL COACTIVATOR 3287
eluted in SDS sample buffer and analyzed by SDS-PAGE and Coomassie bril-
liant blue R-250 (Bio-Rad).
For GST pulldown experiments, glutathione-agarose beads were coated with
bacterial lysates containing GST fusion proteins for 60 min at 4°C and subse-
quently washed three times with H3-binding buffer. Mononucleosomes from
HeLa cells (prepared as described previously [51]) or transfected 293T cell
lysates were mixed with 50 l beads in a total volume of 400 l and incubated
overnight at 4°C in H3-binding buffer. The beads were washed five times with 500
l of H3-binding buffer. Bound proteins were analyzed by immunoblotting.
Transient reporter coactivation assay. For luciferase assays, U2OS cells were
transfected in triplicate. The firefly reporter luciferase construct 5XGal4MLP-
Luc was cotransfected with 50 ng TK promoter-driven Renilla luciferase plasmid
(to normalize for differences in transfection efficiency) and PHF8-pFlag-CMV2
in combination with Gal4 activator constructs as indicated. Cell lysates were
prepared 24 h after transfection, and the luciferase activity was determined using
the Dual-Luciferase Reporter Assay System (Promega).
Coimmunoprecipitation (CoIP). 293T cells were lysed 24 to 40 h after trans-
fection in IP buffer (50 mM Tris-HCl, pH 8, 150 mM KCl, 5 mM MgCl
2
, 0.5 mM
EDTA, 0.1% NP-40, phosphatase, and protease inhibitor cocktails from Sigma
Aldrich). Flag-tagged protein was bound to M2 beads (Sigma-Aldrich) for3hat
4°C, followed by 3 washes with IP buffer. Precipitated proteins were separated by
SDS-PAGE and transferred onto PVDF membranes. The membranes were
developed with the appropriate antibodies and enhanced chemiluminescence
(ECL) (Pierce).
Knockdown experiments and microarray analysis. HeLa cells were trans-
fected with pRetroSuperPuro plasmids containing short hairpin RNAs (shRNAs)
targeting PHF8 (PHF8-shA or -shB) or nontargeting control (NT-sh) (sequences
are given in the supplementary material). One day after transfection, selection
with 2.5 g/ml puromycin was performed for two more days. Cells were har-
vested with Trizol reagent (Invitrogen) for RNA isolation or with nondenaturing
lysis buffer for protein isolation or were fixed and sonicated for qChIP analysis.
PHF8 knockdown was confirmed by quantitative reverse transcription (qRT)-
PCR and immunoblotting.
Three independent knockdown experiments were performed in HeLa cells
with PHF8-shA and control NT-sh. Total RNA (500 ng) was reverse transcribed,
labeled by in vitro transcription with Cy3 and Cy5, and hybridized to whole
human genome arrays in dye swap (Agilent G4112F). After scanning of the
arrays (Agilent G2565BA), features were extracted with GenePix Pro 6.0 and
analyzed with AFM 4.0 (6). Probes with average changes higher than 2-fold were
considered regulated. Nonannotated transcripts, PHF8, and three ambiguous
transcripts (CCDC6, SCD, and CNOT6, which had different probes up- and
downregulated) were excluded from further analysis. For 11 transcripts, regula-
tion was confirmed by qRT-PCR using cDNAs obtained by reverse transcription
of total RNA with Moloney murine leukemia virus (MMLV) enzyme and
oligo(dT)
18
primer. The sequences of qRT-PCR and qChIP primers are listed in
the supplemental material.
Microarray and ChIP-seq data accession number. Data from the ChIP-seq
and microarray experiments were deposited as a SuperSeries under accession
number GSE20753 in the GEO database (http://www.ncbi.nlm.nih.gov/geo/).
RESULTS
PHF8 is a demethylase specific for dimethylated histone H3
lysine 9. Several JmjC-containing proteins have been shown to
specifically demethylate histone H3 methylated at lysine 4, 9,
27, or 36. These enzymes remove methyl moieties by an oxi-
dative reaction that requires an Fe
2⫹
ion and ␣-ketoglutarate
as cofactors. Since the JmjC domain of PHF8 carries all the
amino acids needed for cofactor binding (Fig. 1A), PHF8 was
suggested to be a histone demethylase (25) and was very re-
cently demonstrated to be active in vitro (31). We expressed
full-length PHF8 in Sf9 insect cells using the baculovirus sys-
tem and isolated the recombinant protein by Flag affinity pu-
rification (Fig. 1B). Bulk histones from calf thymus were used
as PHF8 substrates and analyzed by immunoblotting using an-
tibodies specific for different methyl lysine modifications of histones.
We observed a profound reduction of H3K9me2 after incubation
with PHF8. In contrast, we did not detect any change for the
other histone methyl marks, including, H3K9me3, H3K27me3/2/1,
H3K4me3/2, H3K36me3/2, H3K79me3/2, and H4K20me3/2/1 (Fig.
1C). Demethylation was proportional to the amount of enzyme
(see Fig. S1A in the supplemental material) and the reaction
time (see Fig. S1B in the supplemental material) and was
dependent on the cofactors Fe
2⫹
,␣-ketoglutarate, and ascor-
bic acid (see Fig. S1C in the supplemental material). Like
Loenarz and colleagues (31), we also observed weak activity on
H3K9me1 (see Fig. S1B in the supplemental material), but in
contrast, we could not detect significant reduction of
H3K27me2 or H3K36me2 (Fig. 1C). This difference may have
been due to different recombinant proteins used in the two
studies: ours was a full-length PHF8, while theirs was devoid of
the PHD and the C-terminal half. While our work was under
review, other studies were published that also showed
H3K9me2-specific demethylase activity for PHF8 (15, 19, 55).
To verify its demethylase activity in vivo, we expressed a
Flag-tagged truncated version of PHF8 (containing amino ac-
ids 1 to 489 and a C-terminal nuclear localization signal) in
293T cells. After 24 to 48 h, the cells were fixed and subjected
to indirect immunofluorescence with antibodies against Flag
and histone H3 methylated at different lysine residues. The
mean fluorescence signals of 10 of each untransfected and
transfected nucleus were quantified and compared using Stu-
dent’s ttest (see Table S1 in the supplemental material).
H3K9me2 was markedly reduced in cells that overexpressed
wild-type Flag-PHF8(1-489) (Fig. 1D). The JmjC domain con-
ferred the H3K9me2 demethylase activity. This was indicated
by the lost activity of an H247A/D249A mutant predicted to be
defective in cofactor binding (Fig. 1E). Interestingly, the
XLMR point mutant F279S was devoid of activity (Fig. 1F).
This disease mutant displayed aberrant localization to the cy-
toplasm, formed aggregates in vivo (Fig. 1F), and had reduced
solubility at low salt concentrations in vitro (data not shown).
Our observations with this XLMR mutant are consistent with
recent work by Loenarz et al. (31). Intriguingly, we also ob-
served dramatically impaired demethylase activity when we
mutated the putative H3K4me3-binding residues D28 and
W29 to alanine residues in the PHD (Fig. 1G). However, we
could observe a small demethylation activity with this mutant
following prolonged expression (see Table S1 in the supple-
mental material).
While ectopic expression of PHF8(1-489) resulted in a de-
crease in H3K9me1 (Fig. 1H), there were no changes for
H3K9me3 (Fig. 1I), H3K4me2 (Fig. 1J), H3K27me2 (Fig. 1K),
or H3K36me2 (see Table S1 in the supplemental material).
The demethylation of H3K9me1 was deemed to be less robust
both in vivo (Fig. 1H) and in vitro (Fig. S1B). The increased
amounts of H3K9me1 following the demethylation of
H3K9me2 may mask the extent of H3K9me1 activity of PHF8.
Further truncation of the PHF8 protein leading to a protein
spanning amino acids 1 to 352 containing the PHD and JmjC
domain was devoid of demethylation activity in vivo (see Fig.
S2A in the supplemental material). Surprisingly, the full-length
PHF8 was also deemed to be very low in demethylation activity
when expressed in vivo (see Fig. S2B in the supplemental
material). These results suggest that the approximately 100
amino acids following the consensus JmjC domain are impor-
tant for demethylation activity in vivo. Moreover, it is likely
that the C-terminal half of PHF8 exerts a negative effect on the
demethylation activity in vivo, as full-length PHF8 displayed
3288 FORTSCHEGGER ET AL. MOL.CELL.BIOL.
FIG. 1. PHF8 is an H3K9me2 demethylase. (A) Schematic representation of PHF8 with its PHD (red), JmjC domain (cyan), and nuclear
localization signals (magenta). Below, conserved amino acids crucial for metal ion chelation (red) and for binding to H3K4me3 (green) and
␣-ketoglutarate (cyan) are depicted. F279 (blue) is mutated to serine in some patients with XLMR. (B) Recombinant full-length PHF8 was
produced in insect cells and quantified by comparison to a BSA standard after Coomassie staining. The individual lanes were spliced from one gel.
(C) Bulk histones were incubated without (⫺) or with (⫹) recombinant PHF8 and analyzed by immunoblotting using the given specific histone
antibodies. (D to K) Indirect-immunofluorescence analyses of 293T cells transfected for 24 h with Flag-PHF8(1-489)NLS wild type or the
H247A/D249A, F279S, or D28A/W29A mutant. Flag and the indicated histone H3 methyl lysine antibodies were used. Transfected Flag-positive
cells are indicated by the arrowheads. The image field dimension is 68 m. Quantifications of mean fluorescence signals are given in Table S1 in
the supplemental material.
VOL. 30, 2010 PHF8 IS A TRANSCRIPTIONAL COACTIVATOR 3289
diminished activity. Finally, consistent with a published report
(31), overexpression of PHF8(1-489) in HeLa cells did not
yield a change in H3K9me2 (see Fig. S2C in the supplemental
material), suggesting an important role for a cell type specific-
ity factor for demethylation in vivo. Taken together, our results
indicate that PHF8 displays demethylation activity toward
mono- and dimethyl H3K9. However, we found evidence for
cell-type-specific factors regulating the demethylation activity
of PHF8 in vivo. Importantly, we showed the contribution of
the PHD to the enzymatic activity of PHF8 in 293T cells in
vivo, while the domain was shown not to be crucial for its
activity in vitro (31).
The PHD of PHF8 binds histone H3 methylated at lysine 4.
Comparison of the PHF8 PHD with H3K4me3-specific PHDs
of the bromodomain PHD finger transcription factor (BPTF)
and TBP-associated factor 3 (TAF3) (39, 51) revealed conser-
vation of crucial Zn
2⫹
- and H3K4me3-binding residues (indi-
cated in red and green in Fig. 1A). In peptide pulldown assays,
we found that bacterially expressed GST fusion with the PHD
of PHF8 (amino acids 2 to 66) was specifically retained by
biotinylated H3K4me3/2 peptides (amino acids 1 to 17), but
not by H3K4me1, H3K9me3/2/1, and H3K36me3/2/1 (amino
acids 22 to 39) or unmethylated H3 peptides (Fig. 2A). The
PHD of Taf3 was used as a positive control in these assays.
Interaction of the PHF8-PHD with H3K4me3 was not com-
promised by modifications at neighboring residues, such as
methylation of H3R2/H3K9 or phosphorylation of H3T6/
H3S10. In contrast, concomitant phosphorylation of H3T3
blocked K4me3 binding (Fig. 2B). H3T3 phosphorylation is
mediated by the mitotic haspin kinase (11), and this correlates
with the observation that PHF8 dissociates from mitotic chro-
mosomes despite the presence of H3K4me3 (see Fig. S3A and
B in the supplemental material). As expected, mutation of
Zn
2⫹
-binding (H31A/C34A) (data not shown) or aromatic-
cage (D28A/W29A) residues in the PHF8-PHD abolished
H3K4me3 binding (Fig. 2B). Similar results were obtained with
a mammalian expression construct containing both the PHD
and JmjC domain of PHF8 (amino acids 1 to 352). While
wild-type PHF8, the JmjC H247A/D249A mutant, and the
disease mutant (F279S) bound H3K4me3/2 peptides, the PHD
D28A/W29A mutant did not (Fig. 2C).
We next assessed whether the PHD of PHF8 can also inter-
act with nucleosomal H3K4me2/3. Using GST–PHF8-PHD,
but not the D28A/W29A mutant, we could retain mononucleo-
somes derived from HeLa cells (confirmed by immunoblotting
against H2A, H3, and H4), which were enriched in H3K4me3/2
but not H3K9me2 (Fig. 2D). Again GST-Taf3-PHD served as
a positive control. Analysis of PHF8 in vivo using antibodies
against PHF8 revealed colocalization with H3K4me3 and
RNAPII (see Fig. S3A and B in the supplemental material) in
putative euchromatic regions of interphase 293T cells, while it
was depleted at the nuclear periphery and in perinucleolar
regions, where H3K9me2 was enriched (see Fig. S3C in the
supplemental material).
Having established that the PHF8-PHD can bind the
H3K4me3 mark, we examined the observation that PHD in-
tegrity was important for in vivo demethylase activity (Fig. 1G).
To test K4me3 involvement in this, we cotransfected 293T cells
with wild-type Flag-PHF8(1-489)NLS and the H3K4me3-spe-
cific demethylase JARID1A/KDM5A (8). The ability of PHF8
to demethylate H3K9me2 was significantly reduced by con-
comitant H3K4me3 removal (see Fig. S4A in the supplemental
material), while control cotransfections of PHF8 with JMJD3/
KDM6B, an H3K27me3 demethylase (13), did not result in
impaired PHF8 demethylase activity (see Fig. S4B in the sup-
plemental material). The mean fluorescence signals of
H3K9me2/H3K4me3 for PHF8/JARID1A and H3K9me2/
H3K27me3 for PHF8/JMJD3 cotransfections were quantified
FIG. 2. The PHD of PHF8 binds H3K4me3. (A) Bacterial lysates
containing GST-PHF8(2-66) (top) or GST-Taf3(857-924) (bottom)
were incubated with streptavidin beads coated with H3 peptides car-
rying the indicated modifications and a C-terminal lysyl biotin. Bound
proteins were analyzed by SDS-PAGE and Coomassie staining. The
arrowheads indicate the positions of the GST fusion protein. (B) Wild-
type (top) and D28A/W29A mutant (bottom) GST–PHF8-PHD were
analyzed as described for panel A for binding to methylated and
phosphorylated H3 peptides (1 to 17) as indicated above. (C) Lysates
of transfected 293T cells expressing wild-type or mutant Flag-PHF8(1-
352) proteins were incubated with streptavidin beads coated with the
indicated H3 peptides. Bound proteins were analyzed by immunoblot-
ting using Flag antibodies. The arrowheads show the positions of the
indicated proteins, and the asterisks show background binding to H3
peptides. (D) Glutathione beads were coated with GST, GST–PHF8-
PHD, GST–PHF8-PHD mutant (D28A/W29A), and GST-Taf3-PHD
as a positive control and incubated with native HeLa mononucleo-
somes. The bound material was analyzed by immunoblotting using the
given antibodies or Ponceau S staining as indicated.
3290 FORTSCHEGGER ET AL. MOL.CELL.BIOL.
and compared for transfected and untransfected cells (see Fig.
S4C in the supplemental material). This is in agreement with
the stimulation of H3K9me2 demethylation activity on pep-
tides by the presence of H3K4me3 (19). Taken together, these
experiments suggest that the PHD of PHF8 can bind specifi-
cally to H3K4me3-marked nucleosomes and that this binding is
important for its histone demethylase activity in vivo.
PHF8 is a global reader of H3K4me3. In order to determine
the genomic location of PHF8, we performed chromatin im-
munoprecipitation followed by high-throughput sequencing
using the Illumina/Solexa technology (ChIP-seq). Sonicated
chromatin from fixed 293T, HeLa-S3, and serum-starved and
restimulated Hs68 cells was immunoprecipitated with rabbit
anti-PHF8 antibody and normal rabbit IgG as a negative con-
trol. In Hs68 fibroblasts, we used input DNA as a control and
performed ChIP with rabbit PHF8 or H3K4me3 antibody. On
average, 14 million total reads were obtained per flow cell, and
approximately 63% of these could be unambiguously mapped
to the human genome. Cluster analysis of uniquely aligned
reads revealed more than 10,000 significant peaks for PHF8 in
all three cell lines. The ChIP-seq breakdown and cluster iden-
tities are given in Table S2 in the supplemental material. The
peaks were almost exclusively located in close proximity to
TSSs, and we observed strong overlap between PHF8,
H3K4me3, and RNAPII clusters using our own and published
data (37, 38) (Table 1), suggesting that PHF8 preferentially
occupied thousands of active or poised promoters. ChIP-seq
data were validated at several c-myc target loci by quantitative
real-time PCR of chromatin-immunoprecipitated DNA ob-
tained with rabbit PHF8, c-myc, or normal IgG. PHF8 exhib-
ited high (for NCL [see Fig. S5A in the supplemental mate-
rial]), medium (for CDK4,CCNB1, and ITGB1 [see Fig. S5B to
D in the supplemental material]), or no (for TERT [see Fig.
S5E in the supplemental material]) enrichment in these re-
gions, which also corresponded to H3K4me3 and RNAPII
occupancies.
A more detailed strand-specific analysis of sense (5⬘)or
antisense (3⬘) reads in respect to oriented TSSs revealed a
similar distribution and a phase shift of 120 bp for PHF8 in
293T (Fig. 3A) and HeLa-S3 (Fig. 3B) cells and for PHF8 (Fig.
3C) and H3K4me3 (Fig. 3D) in Hs68 cells. No enrichment of
reads for controls, like IgG ChIP of HeLa cells (Fig. 3E), or for
input DNA of serum-stimulated Hs68 cells (Fig. 3F) was ob-
served at TSSs. Active and poised TSSs exhibit defined posi-
tioning of nucleosomes, which tend to be methylated at H3K4
(4, 41). Similar to previously reported experiments using native
ChIP of mononucleosomes (40, 41) or sonicated cross-linked
chromatin (37), we also observed several phased H3K4me3
peaks at the expected nucleosome positions in Hs68 (Fig. 3D).
The phase shift in our experiment was with 120 bp; however, it
was smaller than the 150 bp obtained with mononucleosomes,
possibly because of harsher fragmentation conditions during
the sonication process. Analysis of PHF8 reads indicated one
peak immediately before and one after the TSS (Fig. 3A to C).
Therefore, we concluded that PHF8 bound preferentially to
nucleosomes at the ⫺2 and ⫹1 positions. A short region at the
TSS corresponding to nucleosome position ⫺1 was depleted of
H3K4me3 and PHF8 reads. In active or poised promoters, this
nucleosome position is occupied either by RNAPII and the
basal transcription machinery (41) or by unstable H2A.Z/H3.3-
containing nucleosomes (23). Association of PHF8, H3K4me3,
and PolII in vivo was further substantiated by their colocaliza-
tion in immunofluorescence analysis (see Fig. S3A and B in the
supplemental material).
Next, we determined the overlap of PHF8-occupied genes in
the 3 examined cell lines. Out of 10,876 genes in HeLa, 9,210
in 293T, and 8,014 in Hs68 cells, 6,632 shared PHF8 enrich-
ment at their promoters, while others were occupied in only
one or two of the cell lines (Fig. 3G).
Subsequently, we correlated PHF8 occupancy with expres-
sion and H3K4me3 levels. Basically, all annotated genes occu-
pied by PHF8 in Hs68 cells also carried H3K4me3 peaks, but
we detected about twice as many clusters and occupied genes
for H3K4me3 as for PHF8. We sorted the genes by decreasing
counts of H3K4me3 reads at their promoters, grouped them
into bins of 500, and determined the percentage of PHF8
cooccupancy per bin. While 70% of the promoters with high
and medium H3K4me3 levels were cooccupied by PHF8, less
than 10% of promoters with low H3K4me3 also carried PHF8
(Fig. 3H). Furthermore, the average PHF8 read count at pro-
moters with high H3K4me3 levels was about twice as high as at
those with low levels (data not shown). In a similar analysis, we
took advantage of Affymetrix data for HeLa from a previous
study (10), sorted genes by decreasing expression levels, clas-
sified them into bins of 500, and for each bin determined the
percentage of PHF8 occupancy at their promoters in HeLa
cells. While 90% of genes with high transcript levels displayed
PHF8 occupancy, only 10% of genes with low expression levels
were occupied by PHF8 (Fig. 3I). These results suggest a
positive correlation between genes that are occupied by PHF8
and those that display transcriptional activity. However, about
30% of the genes with high H3K4me3 levels in Hs68 were not
cooccupied by PHF8.
PHF8 is a transcriptional coactivator. PHF8 may activate
transcription, for example, by binding to an activating chroma-
tin mark (H3K4me3) and/or by removing a repressive chroma-
tin modification (H3K9me2). We used a reporter assay to test
these possibilities (51). Cotransfection of human osteosarcoma
TABLE 1. Summary of ChIP-seq cluster analysis
Cell line No. of
PHF8 clusters
% TSS
overlap (⫾1,000 bp)
No. of
occupied promoters
% H3K4me3
overlap (⫾1,000 bp)
% RNAPII
overlap (⫾1,000 bp)
HeLa-S3 19,447 96 10,876 94
a
88
b
293T 13,778 97 9,210 ND
c
ND
Hs68 ⫹FBS 16,411 83 8,014 81 ND
a
Data from Robertson et al. (37).
b
Data from Rozowsky et al. (38).
c
ND, not determined.
VOL. 30, 2010 PHF8 IS A TRANSCRIPTIONAL COACTIVATOR 3291
FIG. 3. PHF8 reads H3K4me3 at TSSs, and occupancy correlates with transcript levels. ChIP-seq reads of PHF8 in 293T (A), HeLa (B), and
serum-stimulated Hs68 (C) cells; of H3K4me3 in serum-starved Hs68 cells (D); of negative-control IgG in HeLa cells (E); and of input in serum-
stimulated Hs68 cells (F) were strand specifically correlated with annotated TSSs in a range from bp ⫺1000 to ⫹1000. Normalized count densities of 5⬘
(red), 3⬘(green), and combined raw (black) reads are shown. The putative positions of phased nucleosomes are numbered and depicted as blue ovals.
(G) Venn diagram depicting the overlap of PHF8-occupied genes in HeLa (blue), 293T (red), and Hs68 (green) cells. (H) Correlation of H3K4me3 and
PHF8 occupancies in Hs68 cells. Genes with H3K4me3 peaks were sorted by read counts and grouped into bins of 500, and the percentage of genes in
the bin that were also occupied by PHF8 was determined. Bins with decreasing H3K4me3 levels are shown from left to right; the corresponding
percentages of PHF8 cooccupancy are given on the yaxis. (I) Correlation of PHF8 and transcript levels in HeLa cells. Genes were sorted by normalized
expression levels and grouped into bins of 500, and the percentage of genes in the bin that were occupied by PHF8 was determined. Bins with decreasing
transcript levels are shown from left to right; the corresponding percentages of PHF8 occupancy are shown on the yaxis.
3292 FORTSCHEGGER ET AL. MOL.CELL.BIOL.
U2OS cells with a firefly luciferase reporter construct and a
construct encoding full-length Flag-tagged PHF8 indicated a
6-fold increase in promoter activity (Fig. 4A). Moreover, ec-
topic expression of PHF8 revealed a coactivator function to-
ward Gal4-tagged Ash2, p53, c-myc, and E2F (Fig. 4A).
Shorter versions of PHF8 (1-352 and 1-489) displayed im-
paired coactivation, although they were expressed at similar
levels, as shown by Western blotting (Fig. 4A). To extend this,
we initially assessed the functional consequences of PHD and
JmjC mutations in the context of truncated PHF8(1-489).
While PHF8(1-489) displayed impaired coactivation capacity
compared with full-length PHF8, it was still able to enhance
E2F activity by about 2.5-fold (Fig. 4B). Interestingly, while the
PHD (D28A/W29A) and XLMR (F279S) mutants displayed
diminished coactivation functions, the JmjC H247A/D249A
mutant showed only a small decrease in coactivator function.
We next tested whether these observations extended to the
full-length PHF8. Similar to PHF8(1-489), in the context of
full-length PHF8, the PHD D28A/W29A mutant or the F279S
mutant was impaired in coactivation of Gal4-p53 or Gal4-Ash2
(Fig. 4C). Again, the catalytic JmjC H247A/D249A mutant
displayed a minor reduction in coactivation. Taken together,
these results indicate that a functional PHD is an important
component of the coactivator function of PHF8 and that the
activity of the JmjC domain may not be an essential determi-
nant, at least in the context of luciferase assays.
PHF8 interacts with the CTD of RNA polymerase II.
Genomic localization analysis of PHF8 revealed a preference
for transcription start sites that are H3K4 methylated and
occupied by RNAPII. We hypothesized that PHF8 could in-
teract with preinitiation complex components like the general
transcription factor TFIID or RNAPII. To examine this, we
performed CoIP experiments with full-length Flag-PHF8 in
293T cells. Immunoblot analysis revealed coprecipitation of the
RNAPII subunits RPB1 (POLR2A) and RPB3 (POLR2C), but not
of the TAF1 subunit of TFIID (Fig. 5A). When we compared
full-length Flag-PHF8 with truncated versions in CoIPs, the
N-terminal half of PHF8 (1-489) displayed a weak association
with RNAPII, and such an interaction could not be detected
with the 1-352 version (Fig. 5B). These results indicate that the
C-terminal half of PHF8 plays an important role in RNAPII
association. This contention is consistent with coimmunoprecipi-
tation results of the full-length PHD D28A/W29A mutant and the
JmjC H247A/D249A mutant, which interacted with RNAPII sim-
ilarly to the wild type (Fig. 5C). Even though larger DNA
amounts were transfected, the XLMR F279S mutant showed
lower levels of expression and consequently reduced amounts of
precipitated RNAPII (Fig. 5C), probably due to mislocalization
and aggregation.
The CTD of the largest RNAPII subunit, RPB1, has been
shown to be a binding platform for many proteins (33). To test
PHF8 interaction with the CTD, we performed GST pulldown
experiments using GST fused to the CTD (consisting of 27
heptad repeats) and lysates from Flag-PHF8-transfected cells.
GST-CTD bound full-length PHF8 and truncated 1-489, but
not 1-352, while GST-Taf3 was used as a negative control for
these experiments (Fig. 5D). Interestingly, none of the
tested PHF8 mutations (D28A/W29A, H247A/D249A, or
F279S) affected association of full-length PHF8 with GST-
CTD (Fig. 5E).
FIG. 4. PHF8 coactivates reporter gene expression. (A) U2OS cells were
transfected in triplicate with the 5XGal4MLP-Luc reporter plasmid and
pcDNA3 empty vector or pFlag-CMV2 expression plasmid for full-length
PHF8 or its truncated forms, 1-352 and 1-489, in the presence or absence of
Gal4-Ash2, Gal4-myc, Gal4-p53, or Gal4-E2F as indicated. The experiment
shown is representative of three biological replicates, with error bars indicat-
ing standard deviations (SD) within the experiment. The graphs represent the
fold activation relative to transfection with the 5XGal4MLP-Luc reporter
plasmid alone. Below, Flag-PHF8 expression was examined by immunoblot-
ting. wt, wild type. (B) U2OS cells were transfected as described for panel A
with the pcDNA3 empty vector or pFlag-CMV2 expression plasmid of
PHF8(1-489) or its D28A/W29A (PHDmut), H247A/D249A (JmjCmut), or
F279S mutant in the presence or absence of Gal4-E2F as indicated.
(C) U2OS cells were transfected as described for panel A with pcDNA3
empty vector or pFlag-CMV2 expression plasmid of full-length PHF8 or its
D28A/W29A (PHDmut), H247A/D249A (JmjCmut), or F279S mutant in
the presence or absence of Gal4-Ash2 or Gal4-p53 as indicated.
VOL. 30, 2010 PHF8 IS A TRANSCRIPTIONAL COACTIVATOR 3293
FIG. 5. PHF8 interacts with the CTD of RNAPII. (A) 293T cells were either mock transfected with pcDNA3 or transfected with pFlag-CMV2
containing full-length PHF8. Cell lysates were collected and subjected to immunoprecipitation using anti-Flag beads (IP M2). Precipitated proteins
were analyzed by immunoblotting with the indicated antibodies. 5%inp., 5% input. (B) 293T cells were either mock transfected with pcDNA3 or
transfected with full-length, 1-489, or 1-352 PHF8. Immunoprecipitation and detection of precipitated proteins was performed as described for
panel A. The RPB1 antibodies 8WG16 and H-224 were directed against the CTD and an N-terminal epitope, respectively. (C) 293T cells were
either mock transfected with pcDNA3 or transfected with pFlag-CMV2 containing full-length PHF8, PHDmut (D28A/W29A), JmjCmut (H247A/
D249A), or F279S mutant. Immunoprecipitation and detection of precipitated proteins were performed as described for panel A. (D) Bacterial
lysates containing GST fusions with the CTD of the RPB1 subunit of RNAPII or with the PHD of murine Taf3 (as a control) were bound to
glutathione-agarose beads. “Empty” indicates that no bacterial lysate was added. The beads were washed and incubated with 293T cell lysates
transfected with full-length PHF8, PHF8(1-489), or PHF8(1-352). Bound proteins were analyzed by immunoblotting using Flag antibodies. The
arrowheads indicate the positions of the PHF8 proteins. (E) GST pulldown experiment with GST fused to the CTD repeats or to UbcH5B (as a
control) using 293T cell lysates transfected with PHF8-pFlag-CMV2 or the PHD (D28A/W29A), JmjC (H247A/D249A), and F279S mutants as
indicated above the lanes. Binding reactions were processed as for panel D. (F) Bacterial lysates containing GST fusions with the CTD repeats
of the RPB1 subunit of RNAPII were bound to glutathione-agarose beads. After being washed, the GST-CTD-coated beads were incubated with
or without CDK7/cyclin H/Mat1 or CDK9/cyclin T as indicated above the blots and ATP as indicated below. The beads were washed and incubated
with 293T cell lysates transfected with PHF8-pFlag-CMV2. Bound proteins were analyzed by immunoblotting using Flag (top) or GST (bottom)
antibodies. The positions of PHF8, GST-CTD, and its phosphorylated form are indicated.
3294 FORTSCHEGGER ET AL. MOL.CELL.BIOL.
Next, we tested whether phosphorylation of the CTD influ-
ences its interaction with PHF8. Phosphorylation of serine 5 in
the heptad repeat is conferred by CDK7 and takes place in the
initiation phase near the transcription start site, while phos-
phorylation of serine 2 by CDK9 is thought to occur in the
transcribed region during processive elongation (33). Follow-
ing preincubation of GST-CTD with the CDK7 or CDK9 ki-
nase, we observed the expected mobility shift in CTD depen-
dent on ATP addition (Fig. 5F). However, phosphorylation of
the CTD using either kinase did not alter the interaction with
PHF8 (Fig. 5F). These results suggest that PHF8 interacts
mainly through its C-terminal half with the CTD of RNAPII
and that this association is not influenced by the phosphoryla-
tion status of the CTD. Taken together, the interaction prop-
erty of PHF8 with RNAPII is consistent with the genomic
localization of these proteins and the transcription coactivator
function of PHF8.
We also examined the possibility that PHF8 may physically
interact with transcriptional activators and coactivators that
were used for functional-coactivation assays (Fig. 4A). To this
end, we performed CoIPs with lysates from 293T cells that
were cotransfected with Flag-PHF8 and Gal4-Ash2, Gal4-p53,
or c-myc. We were unable to detect efficient association of
PHF8 with any of these proteins (see Fig. S6A and B in the
supplemental material). These results indicate that the coacti-
vator function of PHF8 is not mediated by direct association
with transcription factors, but more likely via its interaction
with RNAPII.
Knockdown of PHF8 has specific but not global effects on
gene expression. Next, we investigated the effects of PHF8
knockdown on gene transcription. A specific shRNA (PHF8-
shA) was able to significantly decrease the PHF8 level in HeLa
cells 3 days posttransfection compared to a nontargeting con-
trol (NT-sh), as shown on the RNA level by quantitative RT-
PCR (Fig. 6A) and on the protein level by immunoblotting
(Fig. 6B). Total RNAs from three independent knockdown
FIG. 6. Knockdown of PHF8 in HeLa cells using shRNA affects
specific transcripts. (A) PHF8-shA reduced PHF8 mRNA levels by
about 90% compared with control NT-sh, as determined by quantitative
RT-PCR. PHF8 transcript levels were normalized to ACTB and
GAPDH (glyceraldehyde-3-phosphate dehydrogenase) housekeeping
genes using the 2
⫺⌬⌬CT
method. The values correspond to means ⫾
SD (n⫽3). (B) PHF8-shA reduced PHF8 protein levels compared
with NT-sh. HeLa whole-cell lysates were used for immunoblots using
goat (g) and rabbit (r) PHF8 and mouse (m) -actin antibodies as
indicated. (C) Overview of microarray results. Log
2
values of averaged
normalized fluorescence ratios (PHF8-shA to NT-sh) were plotted
according to their ranks for all 41,000 probes. Probes that were more
than 2-fold changed are indicated by arrows (combined, 1,344 probes,
or 3.3% of the total). (D) Box-and-whisker plot depicting PHF8 read
counts at promoters in untransfected HeLa cells for upregulated and
downregulated gene sets. The numbers of genes that were PHF8 oc-
cupied and their percentages of the annotated genes in the set are
given below. (E) Box-and-whisker plot depicting transcript levels of
untransfected HeLa cells (normalized robust multiarray average
[RMA] values derived from the work of Cuddapah et al. [10]) for the
gene sets that were up- and downregulated upon PHF8 knockdown.
The numbers of genes for which expression data were available and
their percentages of the annotated genes in the set are given below. In
panels C and D, the central lines represent medians, the hinges rep-
resent quartiles, and the whiskers represent minimum and maximum
values. N. S., no significant difference (according to Student’s ttest).
Experiments were performed 72 h after shRNA vector transfection.
VOL. 30, 2010 PHF8 IS A TRANSCRIPTIONAL COACTIVATOR 3295
experiments were compared in two-color microarray dye
swaps; 960 probes were more than 2-fold downregulated, while
384 had over 2-fold-increased levels after knockdown (Fig.
6C). The down- and upregulated probes corresponded to 780
and 270 annotated transcripts, respectively (see Table S3 in the
supplemental material). The observation that almost three
times more transcripts were down- than upregulated is in line
with a coactivator role of the knockdown target. However,
knockdown did not result in a global defect compromising
transcription of all PHF8-occupied genes. Furthermore, the
up- and downregulated genes showed similar percentages of
PHF8 occupancy (about 64%), similar levels of PHF8 reads at
their promoters (Fig. 6D), and comparable transcript levels in
untransfected HeLa cells (Fig. 6E). Functional-annotation
analysis of downregulated transcripts using DAVID (12) re-
vealed enrichment for pleckstrin homology domain-containing
proteins and proteins that are involved in the focal-adhesion
pathway (see Table S3 in the supplemental material). Taken
together, these results indicate that PHF8 does not play a role
in basal transcriptional regulation, as the number of genes
displaying transcriptional responsiveness to PHF8 depletion is
only a fraction (about 5%) of those occupied by PHF8. It is
more likely that PHF8 is required for the fine tuning of the
transcriptional output, which is governed by the signaling path-
ways that regulate the individual PHF8-responsive genes. This
scenario is consistent with our coactivation function for PHF8,
indicating that it could further enhance the activities of diverse
transcriptional activators.
Next, we analyzed the genes that displayed downregulation
following PHF8 knockdown and that have been reported to be
linked to mental retardation. These comprise 11 candidate
genes, i.e., MAOA,DPSYL2,CLN5,AP1S2,SMC1A,IDS,
SLC9A6,CUL4B,OPHN1,OCRL, and AMMECR1. Using
quantitative RT-PCR, we could validate the downregulation of
the candidate genes (see Fig. S7A in the supplemental mate-
rial). While the less effective short hairpin (PHF8-shB) also
resulted in downregulation of these genes, the extent of this
effect was smaller than that seen for the more effective PHF8-
shA (see Fig. S7A in the supplemental material). Moreover,
while we observed a reduction of PHF8 occupancy at the
candidate gene promoters upon PHF8 knockdown, we could
not see a consistent change in H3K9me2 or RNAPII levels (see
Fig. S7B in the supplemental material). Since the family mem-
ber KIAA1718 (also called JHDM1D or KDM7A), which is
able to demethylate H3K9me2 and H3K27me2 (19, 20, 49),
was induced 2.1-fold upon PHF8 knockdown, it might be com-
pensating for the loss of PHF8.
DISCUSSION
The key findings of this study are as follows. First, PHF8 was
delineated as a novel histone demethylase of the JmjC family
specific for dimethyl H3K9 and, to a lesser extent, for the
monomethyl state. Second, the PHD of PHF8 was identified as
an important determinant in recognition of trimethyl H3K4.
Third, a critical function for the PHD in histone demethylation
and transcriptional coactivation was revealed. Fourth, it was
shown that a single point mutant of PHF8 associated with
X-linked mental retardation in humans is defective for histone
demethylation and transcriptional coactivation. Fifth, by ge-
nome-wide location analysis of PHF8, a pattern of promoter
occupancy similar to that of trimethyl H3K4 was revealed.
Finally, a direct association between PHF8 and the CTD of the
large subunit of RNA polymerase II was demonstrated.
We showed that PHF8 is an active H3K9me2 demethylase
similar to the JHDM2/KDM3 family (54). Loenarz et al.
recently additionally reported in vitro demethylation of
H3K27me2 and H3K36me2 using a construct lacking the
PHD and the C-terminal domain of PHF8 (31). However, in
our study, full-length PHF8 was unable to demethylate
H3K27me2 or H3K36me2 and was quite specific for dem-
ethylation of di- and monomethyl H3K9. Moreover, overex-
pression of PHF8(1-489) did not result in a decrease of any
of the investigated histone modifications but H3K9me2 and
H3K9me1 (see Table S1 in the supplemental material). Very
recently, Feng et al. also reported that PHF8(1-690) has only
marginal activity on H3K27- or H3K36-methylated peptides
(15). H3K9me2 is an important repressive chromatin modifi-
cation whose removal signals for coactivation of gene expres-
sion. Indeed the importance of the murine H3K9me2 demeth-
ylase Jhdm2a for proper activation of target genes by removing
this repressive mark at the respective promoters has already
been demonstrated (34, 47). This demethylation may result in
the release of repressive factors, or it may facilitate the recruit-
ment of transcription factors and coactivators.
We found that the PHD of PHF8 plays a critical role in its
demethylation activity in vivo, as well as its transcriptional
coactivation function. Our results indicate that the PHD of
PHF8 serves as a specific reader of H3K4 tri- and dimethyl
modifications and may act as an important determinant in
anchoring PHF8 at transcription start sites of PHF8-occupied
promoters. Moreover, PHF8 occupies a wide spectrum of
H3K4-trimethylated promoters, suggesting that it might con-
tribute to the activating effect of this histone modification on
transcription. This contention is further supported by its ability
to function as a coactivator for a number of transcriptional
activators that we have examined, including Ash2, p53, c-myc,
and E2F. These activators have been linked to recruitment of
the MLL histone methyltransferase complex to mediate H3K4
methylation at promoters (32, 50). Therefore, we propose a
model by which increased H3K4me3 levels, through the action
of an MLL-related family of enzymes, would lead to further
recruitment of PHF8 to the promoter of responsive genes.
Once at the promoter, PHF8, through its association with
RNAPII, may stabilize the preinitiation complex formation,
leading to enhanced transcription (Fig. 7).
We found that the disease-causing mutant of PHF8 (F279S)
displays aberrant cellular localization and is devoid of de-
methylation activity. Furthermore, this mutant exhibited re-
duced activity in our coactivation assays. While we did not
observe a general defect in transcription or increased
H3K9me2 levels upon knockdown of PHF8, a specific tran-
scriptional defect in neuronal cells may underlie the disease
phenotype of this mutant PHF8. Indeed, XLMR patients dis-
play specific defects in the development of neurons and the
midline, arguing against a global role for PHF8 in transcrip-
tion. Possible reasons for this lack of global effects on methyl-
ation levels or transcription following PHF8 mutations could
be that in most tissues, the loss of PHF8 is compensated for by
redundant proteins like JHDM2, PHF2, or KIAA1718, the last
3296 FORTSCHEGGER ET AL. MOL.CELL.BIOL.
of which is induced by PHF8 knockdown, or that the activity
of endogenous PHF8 is strictly regulated by signaling path-
ways and switched on in a tissue-, time-, and/or locus-spe-
cific manner.
Our data suggested that the interaction with H3K4me3 was
crucial for PHF8 function, since mutation of aromatic-cage
residues in the PHD, as well as removal of H3K4me3 by
JARID1A, significantly reduced demethylation and coactiva-
tion. It has been shown that activation of neuron-specific genes
occurs by recruitment of the H3K4 methyltransferase MLL1
during in vitro differentiation of P19 cells (52). On the other
hand, deletions in the H3K9 methyltransferase EHMT1 cause
the 9q34.3 subtelomeric deletion syndrome, one feature of
which is mental retardation (24). Additionally, mutations in
other PHD protein-coding genes, like ATRX and PHF6, are
implicated in X-linked neurological disorders (3). Several
H3K4me3 binders, like the inhibitors of growth protein family
(ING1 to -5), the Taf3 subunit of TFIID, or the NURF subunit
BPTF, have been demonstrated to be involved in gene tran-
scription and chromatin remodeling (42, 51, 53). These pro-
teins display different affinities for the H3K4me3 mark, and
their recruitment to promoters could be stimulated via inter-
action with other transcription factors, DNA, or chromatin.
Another JmjC protein, SMCX/KDM5C, is also implicated in
XLMR, but in contrast to PHF8, one of its two PHDs binds
H3K9me3. SMCX has been shown to act as a demethylase
specific for H3K4me3 and a transcriptional repressor (22, 46).
This argues for the importance of a balanced H3K4 and H3K9
methylation and readout in neuronal development and brain
function (2). Writers, erasers, or readers exhibit synergistic or
opposing roles by interaction at or competition for genomic
binding sites in order to produce the desired transcriptional
outcome. Disturbances in the fine tuning of this delicate equi-
librium brought about by mutations in genes like PHF8 can
result in diseases such as hereditary disorders or cancer.
ACKNOWLEDGMENTS
We thank Kristian Helin, Min Gyu Lee, and Cle´ment Carre´ for
providing reagents and the Ultrasequencing and Microarrays Units at
the CRG for performing Solexa and microarray analyses. We are
particularly grateful to Marc Vigneron (ESBS, Strasbourg, France) for
providing unpublished RNAPII antibodies and the GST-CTD con-
struct. We thank Harm-Jan Vos for help in peptide synthesis, Hetty
van Teeffelen for technical assistance, and Michiel Vermeulen for
sharing unpublished results.
K.F. received an Erwin-Schro¨dinger fellowship from the Austrian
Science Fund FWF (J2728-B12). P.D.G., N.S.O., and H.T.M.T.
were supported by grants from the Netherlands Organization for
Scientific Research (NWO-CW TOP 700.57.302), the European
Union (EUTRACC LSHG-CT-2006-037445), and the Netherlands
Proteomics Center. R.S. was supported by a grant from the NIH
(CA090758).
We declare that we have no competing financial interest.
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