human protein arginine methyltransferases in vivo – distinct ...dynamic modification (bedford,...

667Research Article

IntroductionPost-translational modification of proteins is an important means

to expand the structural and functional diversity of the proteome.

Depending on their function, post-translational modifications can

either be transient or stable. Although transient modifications, such

as phosphorylation or acetylation, form the basis of cellular

signaling and act as reversible activity switches, stable modifications

such as myristylation, glycosylation, or proteolytic processing,

usually affect the physicochemical properties of proteins, often

without direct impact on protein activity. The methylation of

arginine residues appears to be a rather stable, but nonetheless

dynamic modification (Bedford, 2007; Bedford and Richard, 2005;

Fackelmayer, 2005b; Wysocka et al., 2006). It has been implicated

in a large variety of important cellular functions, such as signaling

(Berthet et al., 2002; Blanchet et al., 2006; Mowen et al., 2004; Xu

et al., 2004), DNA repair (for a review, see Lake and Bedford, 2007),

maturation and nucleocytoplasmic transport of RNA (Boisvert et

al., 2002; Cheng et al., 2007; Shen et al., 1998; Yanagida et al.,

2004), protein protection (Fackelmayer, 2005b), ribosomal assembly

(Bachand and Silver, 2004; Swiercz et al., 2007; Swiercz et al.,

2005) and the regulation of gene expression (e.g. An et al., 2004;

Balint et al., 2005; Iberg et al., 2008; Xu et al., 2004). However,

the molecular mechanisms connecting the modification to any

functional consequences are only beginning to emerge. With regard

to the regulation of gene expression, arginine methylation appears

to act as an epigenetic mark on chromatin, similar to lysine

methylation, which contributes to the histone code and affects

expression of the underlying genomic information. In this case,

methylated arginines on histones (reviewed by Ruthenburg et al.,

2007), and possibly non-histone proteins involved in nuclear

architecture (Fackelmayer, 2005a; Herrmann et al., 2004), are

recognized by protein adaptors termed ‘effectors’, which contain

domains that specifically bind to methylated arginine (or lysine)

residues. Well-characterized examples for these domains or

‘modules’ are the chromodomain, tudor domain or the PHD finger

(reviewed by Ruthenburg et al., 2007). These modules recognize

different sets of methylated histones and recruit functional multi-

protein complexes to chromatin to exert the appropriate effect, such

as the activation or repression of a particular gene. Less is known

about the mechanisms by which arginine methylation affects

functions unrelated to gene expression, but it is clear that they also

involve discrimination of methylated versus non-methylated

arginine residues by other proteins, and thereby regulate their

function.

To understand the impact of arginine methylation on cellular

events, efforts to isolate, clone and characterize the enzymes

responsible for this modification have been ongoing in many

laboratories, and led to the discovery of a small, distinct family of

protein arginine methyltransferases (PRMTs). In human cells, the

PRMT family consists of eight canonical members [for an overview,

see Bedford (Bedford, 2007)]. They catalyze the addition of one

or two methyl groups to the guanidino nitrogen atoms of arginine,

resulting in either ω-NG-monomethylarginine, ω-NG,NG-

asymmetric or ω-NG,N�G-symmetric dimethylarginine. PRMTs

have been classified into two groups based on the end product, both

of which first catalyze the formation of monomethylarginine as an

Methylation of arginine residues is a widespread post-

translational modification of proteins catalyzed by a small

family of protein arginine methyltransferases (PRMTs).

Functionally, the modification appears to regulate protein

functions and interactions that affect gene regulation, signalling

and subcellular localization of proteins and nucleic acids. All

members have been, to different degrees, characterized

individually and their implication in cellular processes has been

inferred from characterizing substrates and interactions. Here,

we report the first comprehensive comparison of all eight

canonical members of the human PRMT family with respect

to subcellular localization and dynamics in living cells. We show

that the individual family members differ significantly in their

properties, as well as in their substrate specificities, suggesting

that they fulfil distinctive, non-redundant functions in vivo. In

addition, certain PRMTs display different subcellular

localization in different cell types, implicating cell- and tissue-

specific mechanisms for regulating PRMT functions.

Supplementary material available online at

http://jcs.biologists.org/cgi/content/full/122/5/667/DC1

Key words: Arginine methylation, Subcellular localization and

dynamics, Posttranslational modification, Histone methylation,

Epigenetics

Summary

Human protein arginine methyltransferases in vivo –distinct properties of eight canonical members of thePRMT familyFrank Herrmann1, Peter Pably2, Carmen Eckerich2, Mark T. Bedford3 and Frank O. Fackelmayer2,*1EMBL-CRG Systems Biology Research Unit, Centre for Genomic Regulation (CRG), c/Dr. Aiguader 88, 08003 Barcelona, Spain2Biomedical Research Institute, Foundation for Research and Technology Hellas, 45110 Ioannina, Greece3Department of Carcinogenesis, M. D. Anderson Cancer Center, University of Texas, 1808 Park Road 1C, Smithville, TX 78957, USA*Author for correspondence (e-mail: [email protected])

Accepted 17 November 2008Journal of Cell Science 122, 667-677 Published by The Company of Biologists 2009doi:10.1242/jcs.039933

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668

intermediate. In a second step, the type I enzymes PRMT1 (Lin et

al., 1996), PRMT3 (Tang et al., 1998), CARM1/PRMT4 (Chen et

al., 1999), PRMT6 (Frankel et al., 2002) and PRMT8 (Lee et al.,

2005a; Sayegh et al., 2007) lead to the formation of asymmetric

dimethylarginine, whereas the type II enzyme PRMT5 (Branscombe

et al., 2001; Pollack et al., 1999) and possibly PRMT7 (Lee et al.,

2005b; Miranda et al., 2004) lead to symmetric dimethylarginine.

One more member of the PRMT family, PRMT2, was identified

by sequence homology to the known enzymes (Katsanis et al.,

1997), but has not been classified into a group, because no

enzymatic activity has yet been demonstrated or substrates

identified; nevertheless, PRMT2 acts as a coactivator in hormone-

dependent gene expression (Meyer et al., 2007; Qi et al., 2002).

Recently, an additional arginine methyltransferase activity has been

described for FBXO11 (Cook et al., 2006). Unlike the canonical

PRMTs, FBOX11 is not a member of the seven-β-strand

methyltransferase family. In addition, no methyltransferase activity

was reported for the human or worm FBOX11 proteins (Fielenbach

et al., 2007). Consequently, further work is needed to determine

whether FBXO11 is a PRMT, and we have not studied it the context

of this article. In plants, PRMT10 and PRMT11 have been described,

which are true members of the PRMT family, but have no known

human homologs (Niu et al., 2007; Scebba et al., 2007).

Available data suggest that most PRMTs are housekeeping

enzymes, which are ubiquitously expressed in most – if not all –

cell types and tissues of the human body. The only known exception

is PRMT8, which appears to be restricted to neurons in the brain

(Lee et al., 2005a; Sayegh et al., 2007; Taneda et al., 2007).

Importantly, PRMTs differ in their substrate specificities, and are

therefore probably involved in different physiological processes (for

a review, see Bedford, 2007). This has clearly been shown for the

two best-characterized family members, PRMT1 and

CARM1/PRMT4, where the genetic knockout of either enzyme

results in an accumulation of hypomethylated substrates in vivo,

demonstrating that these two PRMTs cannot substitute for each other

(Kim et al., 2004; Pawlak et al., 2002). The activity of both enzymes

is essential for distinct differentiation processes, but is apparently

dispensable for basic cellular metabolism and survival (Pawlak et

al., 2000; Yadav et al., 2003). In this context, it is interesting to

note that the two recently identified PRMTs in plants, PRMT10

and PRMT11, are also required for differentiation processes such

as flowering, but are dispensable for basic cellular life (Niu et al.,

2007; Scebba et al., 2007).

Given the important role of arginine methylation, surprisingly

little is known about the cell biology of the PRMT enzymes. In

particular, data is scarce with respect to subcellular localization,

dynamics and complex formation of PRMTs, which might be central

determinants of their regulation and function in vivo. We have

recently demonstrated that a variant of PRMT1 is primarily localized

in the cytoplasm, spatially separated from the main pool of its

substrates, hnRNP proteins and core histones in the nucleus

(Herrmann et al., 2005). Importantly, these experiments also

demonstrated that PRMT1 is a highly dynamic enzyme with

variable subcellular localization and mobility, dependent on the

methylation status of its substrate proteins. In ongoing efforts to

determine the physiological role of the individual members of the

PRMT family, we report here on the first comprehensive comparison

of subcellular localization and dynamics of all eight canonical

PRMT family members in human cells. Our results provide evidence

that the individual members of the family differ significantly in

their in vivo properties, compatible with distinct physiological

functions.

Journal of Cell Science 122 (5)

Fig. 1. Expression and activity of human PRMT1-PRMT8 fused to EGFP. (A) Schematic representation of the eight canonical members of the human PRMTfamily. The highly conserved PRMT core region (gray), signature motifs I, post I, II, and III (black) and the conserved THW loop (red), present in all PRMTs, areindicated. Note that PRMT7 has a duplication of these motifs and PRMT2 and PRMT3 have an N-terminal SH3 domain (orange) or a zinc-finger domain (green),respectively. The nuclear localization signal (NLS, blue) targets PRMT6 to the nucleus, whereas N-terminal myristoylation (~) tethers PRMT8 to the plasmamembrane. The size of individual PRMTs is indicated at each C-terminus. (B) Human embryonic kidney (HEK293) cells were stably transfected with expressionvectors encoding PRMT1-GFP to PRMT8-GFP. Total cell extracts were separated by SDS-PAGE, blotted to a polyvinylidene difluoride membrane, and probedwith antibodies specific for GFP. Equal amounts of total cell extract were used in each lane, verified by using hnRNP-C as a loading control. (C) Methylationactivity assays. Individual PRMTs were immunoprecipitated from the total cell extracts described in B, by using anti-GFP antibodies. Activity assays were carriedout with hypomethylated extracts (upper panel) or recombinant core histones (lower panel) as described in the Materials and Methods.

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669Arginine methyltransferases in vivo

ResultsThe methylation of arginine residues in proteins has been intensively

studied over the past few years. Many substrates, as well as the

family of enzymes that catalyze the methylation reaction, have been

identified, revealing that arginine methylation is an important

regulator of protein function in a variety of fundamental cellular

processes. However, little is known about the properties of protein

arginine methyltransferases in live cells. In addition, up to now,

studies usually focussed on the properties and functional roles of

individual members of the PRMT family. We therefore decided to

compare, in a comprehensive way, key properties of all eight

canonical PRMT family members under identical conditions. As a

first step towards in vivo measurements, we created human

embryonic kidney (HEK293) cell lines stably expressing GFP fusion

proteins for all eight members of the PRMT family (Fig. 1A). After

four to six weeks of G418 selection, populations containing

expressing cells were obtained by fluorescence-activated cell sorting

(FACS). To ensure that the GFP fusion proteins behave like their

endogenous counterparts and can be used with confidence to

investigate their properties, expressed proteins where checked by

a variety of biochemical means (see Herrmann et al., 2005). All

eight PRMT-GFP fusion proteins were well expressed, and migrated

at their expected molecular mass on SDS gels with no apparent

degradation (Fig. 1B). The expression level of the individual stable

cell lines differed noticeably, with PRMT3, PRMT5, PRMT7 and

PRMT8 being weaker than PRMT1, PRMT2, PRMT4 and PRMT6.

As all eight proteins were expressed equally well in transient

transfections, we assume that the different levels of expression in

stable lines indicate that elevated levels of PRMT3, PRMT5,

PRMT7 and PRMT8 lead to a growth disadvantage, and were thus

overgrown by weaker expressing cells. To measure enzymatic

activity, we immunoprecipitated all constructs by using an anti-GFP

antibody, and used them together with hypomethylated total cell

extract (Fig. 1C, upper panel) or recombinant histones H3 and H4

(Fig. 1C, lower panel) as substrates in radioactive methylation

assays. As expected, strong activity was observed for PRMT1,

followed by PRMT4, PRMT6 and PRMT8. Weak activity, using

total cell extract as a substrate, was seen for PRMT5 and PRMT2,

whereas PRMT3 and PRMT7 did not show detectable activity in

these assays. This result was not surprising, because both enzymes

are known to have very low specific activity (Miranda et al., 2004;

Tang et al., 1998).

Because the formation of homo-oligomeric complexes is a

known prerequisite for enzymatic activity of several PRMTs

(Higashimoto et al., 2007; Lim et al., 2005; Zhang and Cheng, 2003;

Zhang et al., 2000), we determined the migration of our PRMT

constructs on glycerol gradients, for comparison with the migration

of the endogenous proteins (Fig. 2). Extracts from cells stably

expressing PRMT1 to PRMT7 were sedimented in glycerol

gradients; an extract from non-transfected cells was sedimented in

parallel. PRMT8 was omitted from this experiment, because it binds

to the plasma membrane (Lee et al., 2005a) and cannot be extracted

in a membrane-free, soluble form. Fractions from the gradients were

collected and probed with anti-GFP antibodies for the PRMT

constructs, or with commercial antibodies recognizing endogenous

PRMT1 to PRMT7; these experiments were performed for all

PRMTs separately (not shown), and, with identical results, with a

mixture of extracts from the seven cell lines expressing the proteins,

to allow for a direct comparison and to rule out potential gradient-

to-gradient variability (shown in Fig. 2, upper panel). We found

that all GFP fusion proteins, with the exception of PRMT5 (see

below), sediment in identical or very similar way to the endogenous

proteins. PRMT3, PRMT6 and PRMT7 sediment in the gradient

as expected for a monomeric protein or a small oligomer, whereas

PRMT1 and PRMT4, which are known to form homo-oligomers

and associate with a variety of other proteins, sediment as high

molecular mass complexes. These results indicate that all GFP

fusion proteins, except for PRMT5, are properly folded and

oligomerize in their correct functional context. Thus, they are

suitable tools to investigate the localization and dynamics of

PRMTs in vivo. However, PRMT5-GFP did not behave in a similar

manner to the endogenous counterpart; it sedimented as if it was

part of a very large heterogenous complex, suggesting that it is prone

to aggregation (see supplementary material Fig. S2). In vivo

mobility measurements for PRMT5 are shown for completeness

only (see below), but do not represent the behavior of the

endogenous enzyme.

Localization of PRMT-GFP fusion proteins in live cellsThe subcellular localization of members of the PRMT family has

been described previously on an individual basis, but often results

even for the same PRMT are quite variable (Cote et al., 2003;

Frankel et al., 2002; Herrmann et al., 2005; Tang et al., 1998). In

the case of PRMT1, most inconsistencies have recently become

comprehensible, when Goulet and colleagues reported that PRMT1

isoforms produced by alternative splicing localize differently

(Goulet et al., 2007); for our experiments, splicing variant 2 was

used, designated PRMT1v2, which has an N-terminal nuclear export

signal. Our cell lines stably expressing GFP fusion proteins were

an ideal tool to compare the localization of all family members under

identical conditions in live cells (Fig. 3). We found that PRMT1v2,

Fig. 2. Determination of complex sizes of PRMT1-PRMT7 by glycerolgradient centrifugation. A mixture of total cell extracts from cells stablyexpressing PRMT1-GFP to PRMT7-GFP fusion proteins was loaded on a 10-30% glycerol gradient and proteins were separated by centrifugation for 20hours at 30,000 r.p.m. The gradient was fractionated from the top, and aliquotsof individual fractions were resolved by SDS-PAGE. Proteins were detectedby western blotting using anti-GFP antibodies for detection (top panel).PRMT-GFP fusion proteins are denoted by their respective numbers (1-7). Thesedimentation of endogenous PRMTs was investigated similarly, using extractsfrom nontransfected cells and commercially available antibodies for PRMT1to PRMT7. BSA (66 kDa, 4.2S), β-amylase (200 kDa, 8.9S) and apoferritin(443 kDa, 17.6S) were used as sedimentation markers in an identical gradientrun in parallel. Note that the expressed GFP fusion proteins of all PRMTsexcept PRMT5 sediment similarly to their endogenous counterparts.

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PRMT2, PRMT3, PRMT4 and PRMT7 were found both in the

nucleus and in the cytoplasm, ranging from a predominant nuclear

(PRMT2) to a predominantly cytoplasmic (PRMT1v2, PRMT3,

PRMT4 and PRMT7) localization. PRMT6 is exclusively localized

in the nucleus, whereas PRMT5 is only cytoplasmic and PRMT8

is tethered to the plasma membrane. PRMT2 was found

predominantly in the nucleus, and was enriched in subnuclear

structures resembling nuclear speckles (supplementary material Fig.

S3A). In most cases, these localizations were in line with earlier

reports (Frankel et al., 2002; Lee et al., 2005a; Meyer et al., 2007;

Rho et al., 2001; Tang et al., 1998), but especially in the case of

PRMT1 and CARM1/PRMT4 there was some discrepancy that we

wanted to investigate further. We hypothesized that the localization

could be cell-type specific, and created stable cell lines for

PRMT1v2 and CARM1/PRMT4 from often-used parental cell lines

U2OS, HeLa and MCF7. After identical selection and cell sorting,

the cell lines were investigated for localization of the fusion

proteins (Fig. 4). Indeed, the localization of both PRMT1v2 and

CARM1/PRMT4 differed fundamentally in different cell types.

Interestingly, they also did it in an identical way: in U2OS

osteosarcoma cells, PRMT1v2 and PRMT4 were predominantly

cytoplasmic, as in the original HEK293 embryonic kidney cells,

but some cells displayed enrichment in the nucleus. Further

investigations suggested that these differences are due to a cell-

cycle-dependent relocation, especially in the case of PRMT4

(supplementary material Fig. S4). By contrast, in stably transfected

HeLa cervix carcinoma and MCF7 breast cancer cell lines, both

PRMT1v2 and CARM1/PRMT4 were predominantly nuclear. We

conclude that the localization of the PRMTs is cell-type specific,

and can also dynamically change in the cell cycle or under certain

conditions (see Herrmann et al., 2005).

Dynamics of PRMT family members in vivoTo gain qualitative insight and quantitative data on the intracellular

dynamics of the PRMT family members, we performed

photobleaching experiments (FRAP, fluorescence recovery after

photobleaching) as described earlier for PRMT1 (Herrmann et al.,

2005). For each member of the PRMT family we measured 15 to

20 individual cells, using a rectangular bleaching area encompassing

both the cytoplasm and the nucleus; typical examples of the

bleaching and the recovery over time are given in Fig. 5. Pixel

intensities in the bleached region (and, for reference and

normalization, in an unbleached region of the same cell) were

determined, and graphically represented (Fig. 6) to show the


Fig. 3. Subcellular distribution of PRMTsin HEK293 cells. Confocal slices throughtypical cells that stably express PRMT1-GFP to PRMT8-GFP fusion proteins. Forreference, the same cells were analyzedby differential interference contrast (DIC).Scale bar: 25 μm.

Fig. 4. Subcellular distribution of PRMT1 and PRMT4 show cell-typespecificity. Cell lines stably expressing PRMT1-GFP or PRMT4-GFP werecreated from HEK293 (human embronic kidney), U2OS (humanosteosarcoma), HeLa (human cervix carcinoma) and MCF-7 (human breastadenocarcinoma) cells. Fields of typical cells for each construct and cell lineare shown. Scale bar: 50 μm.

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kinetics of recovery. Furthermore, data from the recovery were fitted

to a modified exponential association function (supplementary

material Fig. S1) to determine the half-time of fluorescence recovery

(t/2) and mobile fraction for all PRMTs. The recovery half-time

allowed calculation of the diffusion coefficient for every PRMT

(Table 1A), separately for the nucleus and the cytoplasm, or, in the

case of PRMT8, on the plasma membrane.

Mobile fractions of PRMT1 to PRMT7 were between 94% and

98%, comparable with GFP alone, demonstrating that the enzymes

behave as completely soluble (i.e. not structurally bound) proteins

in both the cytoplasm and the nucleus. For PRMT8, the mobile

fraction in the membrane was 74%, in line with other membrane

proteins (Bates et al., 2006). Interestingly, the mobility of individual

members of the PRMT family was vastly different (Table 1). The

most mobile enzymes were PRMT3 and PRMT7, with recovery

half-times of 0.97±0.08 and 0.58±0.07 seconds in the cytoplasm

and 0.95±0.11 and 1.04±0.08 seconds in the nucleus, respectively.

Only these two members of the PRMT family behave, in live cells,

like monomers or small oligomers when compared with a control

protein, a tetrameric β-galactosidase-GFP construct with a

molecular mass of approximately 600 kDa. Of the remaining

PRMTs, which all show diffusion characteristics of high molecular

weight complexes in these measurements, PRMT1 and PRMT4

(and PRMT5, but these data do not reflect the endogenous protein,

see above) were least mobile in the cytoplasm, and PRMT1 and

PRMT6 were least mobile in the nucleus. For all PRMTs that have

both a cytoplasmic and a nuclear population, the nuclear population

was less mobile than the cytoplasmic population, as expected,

because of the known higher intranuclear viscosity (Wachsmuth

et al., 2000) (and unpublished measurements from our laboratory).

The only notable exception was PRMT3, which did not differ

significantly between nucleus and cytoplasm, indicating that

PRMT3 is slowed down by transient interactions with cytoplasmic

proteins.

Identical in vivo mobility assays were repeated in cells that had

been incubated with the methylation inhibitor periodate-oxidized

adenosine (Adox) (Fig. 6; Table 1B), because we had recently found

that this treatment leads to an immobile fraction of 26% of PRMT1

in the nucleus (Herrmann et al., 2005). This effect was reproduced

here. In addition, the nuclear but not the cytoplasmic fraction of

PRMT3 showed a similar effect, although to a weaker extent,

leading to 12% immobile PRMT3 upon Adox treatment. The mobile

fractions of all other PRMT family members were not affected by

Adox treatment. Control experiments published earlier, using

hnRNP-C and a β-galactosidase model protein showed that Adox

treatment does not measurably affect intranuclear viscosity

(Herrmann et al., 2005). We found, however, that diffusion of several

PRMTs was considerably altered by Adox treatment (Table 1B). In

particular, diffusion of nuclear PRMT2, PRMT3 and

CARM1/PRMT4 was significantly (P<0.001) reduced, whereas a

subtle, yet statistically significant (P<0.001), increase in the

diffusion of nuclear PRMT6 was observed. In the cytoplasm, the

mobilities of PRMT2, PRMT3 and PRMT7 were also decreased

(P<0.001). For PRMT1, the difference was not significant, neither

in the nucleus (P=0.055) nor in the cytoplasm (P=0.019). Also, no

significant difference was found for cytoplasmic PRMT4 (P=0.685)

and cytoplasmic PRMT5 (P=0.012). These results show that PRMT

family members react differently to the accumulation of

unmethylated substrates, and also that individual PRMTs are

differently affected in different subcellular locations, probably

Fig. 5. Photobleaching experiments reveal invivo dynamics of PRMTs. Selected images fromconfocal FRAP experiments on HEK293 cellsthat stably express PRMT1-GFP to PRMT8-GFP. Typical cells are shown before and directlyafter photobleaching, and at distinct time pointsduring fluorescence recovery. Bleaching wasperformed using a rectangular region spanningboth the cytoplasm and the nucleus.

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reflecting the presence of cognate substrates with which transient

interactions can occur.

PRMT complex size determined by FCSIn vivo mobility measurement of the eight PRMT family members

demonstrated that, except for PRMT3 and PRMT7, all PRMTs have

diffusion characteristics of multiprotein complexes. In these

experiments, mobility is affected by many factors, including

transient interactions with substrates or other binding partners, size

and shape of the enzymatically active oligomer, and intracellular

viscosities. To complement the in vivo measurements, we performed

fluorescence correlation spectroscopy (FCS) measurements for all

PRMTs in diluted ex-vivo cell extracts. Under these conditions, the

effects of medium viscosity and transient interactions should be

minimized or absent, allowing for measurement of diffusion

characteristics of the enzymes in stable complexes. To this end,

extracts from cells expressing individual PRMT-GFP fusion proteins

were prepared, and diluted with isotonic buffer to 1-5 molecules

per confocal volume of measurement. Identical experiments were

performed with extracts from untreated cells and cells treated with

Adox. Auto-correlation curves were obtained (Fig. 7) and quantified

(Table 2) for PRMT1 to PRMT7, and for GFP alone as a calibration

standard. All auto-correlation curves could be successfully fitted

with a one-component algorithm, directly yielding a value for τd,

which represents the characteristic diffusion time of the molecules

through the measuring space. As described in the Materials and

Methods, τd was used to determine the diffusion coefficient, and

to estimate the hydrodynamic radius and the apparent molecular

mass of the measured enzymes. Results show that Adox treatment

does not affect the diffusion of any PRMT under these conditions

(Table 2A), revealing that the effects of Adox on in vivo mobility

shown in Table 1B do not reflect changes in the stable enzyme

complexes, but rather in their transient interaction with substrates.

These results are also in excellent agreement with those from

glycerol gradient centrifugation and in vivo mobility measurements

in untreated (non-inhibited) cells. They show that PRMT1 and

PRMT4 are present in high molecular mass complexes, and that

PRMT5-GFP forms large, presumably non-physiological

aggregates. By contrast, FCS data for PRMT2 are compatible with

the presence of a dimer, and data for PRMT3 suggest the existence

of a tetramer (see Discussion). Clearly, PRMT6 and PRMT7 both

behave as monomeric proteins under these conditions.

Taken together, our results provide the first comprehensive

comparison of all eight canonical human PRMTs with regard to

localization, intracellular mobility in live cells and the size of the

active complexes. They show that the enzymes all differ in their

properties, and therefore apparently perform distinct, non-redundant

functions in the cell.

DiscussionThe family of protein arginine methyltransferases, with their eight

canonical members in humans, has been functionally implicated in

a wide variety of cellular processes. Here, we performed the first

full-family comparison of the PRMT enzymes, because a profound

and quantitative knowledge of their properties is essential to

understand the functional diversity in the PRMT family.

We show that GFP fusion proteins can be used as in vivo tools

for studies of intracellular localization and mobility of individual

members of the PRMT family. Except for PRMT5-GFP, which

appears to form large (presumably non-physiological) aggregates,

fusions of the seven other members of the PRMT family behave

like their endogenous counterparts. Thus, although we cannot

completely rule out the idea that the GFP tag affects some properties

of the tagged enzymes, such as the substrate specificity (Pawlak et

al., 2002), we are confident that the fusion proteins well reflect the


Fig. 6. Quantitative analysis of FRAPexperiments. Mobility of PRMTs was quantifiedindividually for the nuclear (A) and thecytoplasmic (B) fraction of the proteins with orwithout oxidized adenosine (Adox) treatment.Note that PRMT5 lacks a nuclear fraction, andPRMT6 lacks a cytoplasmic fraction. Therecovery curve of membrane bound PRMT8 isshown separately (C). Each recovery curverepresents the mean of 12-22 individual cells.

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behavior of the endogenous proteins. Microscopic live-cell analysis

demonstrated that PRMTs differ in their subcellular localization,

mostly but not entirely in line with earlier data obtained for the

individual PRMTs. For example, PRMT6 was found exclusively in

the cell nucleus, owing to its nuclear localization signal (Frankel

et al., 2002; Herrmann et al., 2005) and PRMT8 was found at the

plasma membrane as a result of its N-terminal myristoylation (Lee

et al., 2005a). PRMT5 was the only PRMT that we found to be

completely cytoplasmic in our cells. This was compatible with

earlier data from immunofluorescence experiments (Rho et al.,

2001), but at variance with studies that also show functions for

PRMT5 in the nucleus (Ancelin et al., 2006; Lacroix et al., 2008;

Pal et al., 2003). As noted above, however, we assume that

PRMT5-GFP is aggregated (see supplementary material Fig. S2)

and therefore does not reflect the behavior of the endogenous

protein. The remaining PRMT1, PRMT2, PRMT3, PRMT4 and

PRMT7 are found in the cytoplasm as well as in the nucleus,

although the relative amounts in these compartments differ

significantly. The localization of PRMT1 in our experiments was

especially interesting, because the laboratory of Jocelyn Coté had

recently identified a functional nuclear export signal in the N-

terminus of the splicing variant used in this article (Goulet et al.,

2007). In fact, splicing variant 2 of PRMT1 (PRMT1v2), was

localized predominantly in the cytoplasm in the HEK293 cells we

originally investigated. This was also in line with earlier data from

our laboratory showing that PRMT1 is approximately six- to

eightfold more abundant in the cytoplasm than in the nucleus, but

that this ratio is variable in response to the methylation status of

the cell (Herrmann et al., 2005). More recently, we found that a

catalytically inactive PRMT1 accumulates further in the nucleus

(Herrmann and Fackelmayer, 2009). This suggests that the presence

of a nuclear export signal alone is not sufficient to completely

understand the behavior of PRMT1v2. In fact, the situation is more

complex, because cells from other cell types revealed striking

differences in the localization of PRMT1v2, after creating stable

cell lines with the identical expression construct. We found that,

despite the presence of the nuclear export signal, PRMT1v2 is

predominantly nuclear in stably transfected HeLa and MCF-7 cells,

and in a fraction of U2OS cells. This is compatible with cell-type-

specific regulation of the intracellular shuttling of PRMT1 in and

out of the nucleus. It is interesting to note that CARM1/PRMT4

behaves in a similar way, being a predominantly cytoplasmic protein

in HEK293 cells, but a mainly nuclear protein in MCF-7 and HeLa

cells. More work will be necessary to investigate this shuttling in

detail, because it might provide a better understanding of the

regulation of substrate proteins, and might link to pathophysiological

consequences of its misregulation (Goulet et al., 2007). When the

intracellular mobility of PRMT1 and CARM1/PRMT4 was

investigated by FRAP analysis, we found that both proteins showed

diffusion characteristics of large multiprotein complexes. These

complexes probably consist of multiple copies of the same enzyme,

which has previously been found to be a prerequisite for enzymatic

activity (Higashimoto et al., 2007; Lim et al., 2005; Zhang and

Cheng, 2003; Zhang et al., 2000), and of transiently interacting

proteins that might act either as substrates or regulators of activity

(Cote et al., 2003; Herrmann et al., 2004; Lin et al., 1996; Xu et

al., 2004). Photobleaching experiments show that complexes

containing PRMT1 or CARM1/PRMT4 differ in their mobility, both

in the nucleus and the cytoplasm, suggesting that at least the bulk

of the two enzymes do not interact with each other. This is most

pronounced when methylation of substrates is inhibited. Under these

conditions, PRMT1 gains an immobile fraction in the nucleus of

live cells because it is tightly tethered to unmethylated substrates

(see Herrmann et al., 2005), but PRMT4 is not affected. In addition,

Table 1. Quantitative diffusion analysis of PRMT photobleaching experiment

Cytoplasm t/2 ± s.e.m. Nucleus t/2 ± s.e.m. Cytoplasm D ± s.e.m. Nucleus D ± s.e.m.Protein n (seconds) (seconds) (µm2/second) (µm2/second)

A. Analysis in untreated HEK293 cellsPRMT1 20 2.75±0.21 4.61±0.42 1.83±0.12 1.20±0.12PRMT2 17 0.97±0.08 3.51±0.33 5.28±0.44 1.56±0.16PRMT3 15 0.90±0.08 0.95±0.11 5.53±0.44 5.55±0.52PRMT4 15 1.61±0.13 2.63±0.29 3.12±0.21 1.98±0.17PRMT5 12 3.59±0.36 n.d. 1.44±0.15 –PRMT6 16 n.d. 3.94±0.13 – 1.18±0.04PRMT7 20 0.58±0.07 1.04±0.08 9.21±0.64 4.99±0.38PRMT8 16 24.78±2.09* n.d. 0.21±0.03* n.d.EGFP 8 0.23±0.07 0.56±0.08 22.18±3.14 9.59±1.20βGal-NLS 19 n.d. 2.68±0.14 – 1.80±0.09

B. Analysis in HEK293 cells treated with 15 μM oxidized adenosine for 48 hoursPRMT1 24 2.61±0.16 4.01±0.29 1.93±0.13 1.26±0.07PRMT2 21 1.67±0.10 5.35±0.35 2.98±0.19 0.92±0.05PRMT3 13 1.28±0.11 2.10±0.22 3.86±0.27 2.72±0.36PRMT4 18 1.61±0.12 4.69±0.42 3.09±0.21 1.14±0.11PRMT5 17 3.32±0.39 n.d. 1.58±0.12 –PRMT6 24 n.d. 3.10±0.17 – 1.59±0.09PRMT7 11 0.65±0.07 0.97±0.11 7.82±0.64 5.82±0.74PRMT8 – n.d. n.d. n.d. n.d.EGFP – n.d. n.d. n.d. n.d.βGal-NLS 19 n.d. 2.46±0.11 – 1.94±0.09

Recovery curves shown in Fig. 6 were fitted to a modified exponential association function as described in Materials and Methods, to allow determination ofthe half-time (t/2) of fluorescence recovery. Diffusion coefficients D were calculated from t/2 as described in Materials and Methods. t/2 and D are given as mean± s.e.m. separately for cytoplasmic and nuclear fractions of individual PRMTs. PRMT8 was measured at the plasma membrane; the value is given in the‘Cytoplasm’ column for easier arrangement, and is marked with an asterisk. Between 12 and 22 cells (n) were analyzed for each PRMT. Monomeric EGFP (28kDa) and tetrameric β-galactosidase–EGFP (600 kDa) were analyzed for reference.

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glycerol gradient centrifugation (Fig. 2) shows that, although both

enzymes sediment in overlapping fractions, the peaks are found in

different fractions, with PRMT4 in slightly smaller complexes.

Fluorescent correlation spectroscopy in ex vivo extracts confirms

these different complex sizes, although the differences are not

statistically significant. Interestingly, immunoprecipitation

experiments suggested a possible interaction between PRMT1 and

CARM1/PRMT4, because they could be co-immunoprecipitated to

a certain degree (data not shown). It is conceivable that these

enzymes can join forces in large multiprotein complexes to provide

methylation activity with non-overlapping substrate specificity

under some conditions. This would explain the finding that PRMT1

and CARM1/PRMT4 cooperate in transcriptional activation (An et

al., 2004; Kleinschmidt et al., 2008), and that both are necessary

to affect target gene expression.

PRMT2 was predominantly found in the nucleus, where it forms

a granular pattern and displays regions of enrichment that resemble

nuclear speckles. Ongoing work in our laboratory is investigating

the potential role of PRMT2 in RNA maturation; we would like to

note that we have also, for the first time, found weak enzymatic

activity of PRMT2 (supplementary material Fig. S3C).

Photobleaching revealed that PRMT2 diffuses more slowly in the

nucleus than expected for its size and diffusion characteristics in

vitro, where it behaves like a dimer. This suggests that it forms

transient interactions with nuclear proteins that reduce its mobility.

A similar effect was observed for PRMT6, which sediments as a

free protein and behaves like a monomer in fluorescence correlation

spectroscopy, but displays an intranuclear mobility between those

of the multiprotein complexes containing PRMT1 or

CARM1/PRMT4. Interestingly, inhibition of methylation by Adox

slightly increases the mobility of PRMT6, in contrast to all other

PRMTs that were either unaffected within the limits of statistical

significance (PRMT1 and 7), or decreased (PRMT2,

CARM1/PRMT4, and most significantly, PRMT3). PRMT3 is

known to methylate both nuclear and cytoplasmic proteins, such

as the nuclear poly(A)-binding protein (PABPN1) (Fronz et al.,

2008; Smith et al., 1999) and, most prominently, the cytoplasmic

ribosomal protein S2 (rpS2) (Bachand and Silver, 2004; Choi et

al., 2008; Swiercz et al., 2007; Swiercz et al., 2005). The localization

in both compartments was therefore not a surprise. However,


Fig. 7. Diffusion characteristics of PRMT complexes investigated byfluorescence correlation spectroscopy. Comparison of normalized autocorrelation functions of diluted total cell extracts from cells stably expressingGFP fusion proteins. Cells were treated with oxidized adenosine (Adox) (B) 2days prior to harvesting, or left untreated (A). Note that PRMT1, PRMT4 andPRMT5 show diffusion characteristics of high molecular weight complexes.No significant difference between auto correlation functions in –Adox and+Adox samples was observed.

Table 2. Quantitative diffusion analysis of fluorescence correlation spectroscopy experiments

– Adox + AdoxProtein τ ± s.e.m. (μseconds) τ ± s.e.m. (μseconds)

A. Comparison of diffusion times (t) in extracts from untreated or Adox-treated cellsPRMT1 272.6±6.5 278.4±14.7PRMT2 122.3±3.2 137.0±1.3PRMT3 167.8±6.2 159.3±7.0PRMT4 261.0±3.0 267.3±8.6PRMT5 641.2±33.5 710.0±14.0PRMT6 100.1±0.6 121.0±4.2PRMT7 116.8±2.5 108.5±2.0EGFP 70.3±0.9 n.d.

Protein τ ± s.e.m. (μseconds) D ± s.e.m. (μm2/second) rH ± s.e.m. (nm) Approx. mass ± s.e.m. (kDa) Mass monomer (kDa)

B. Measured parameters for each PRMT in untreated cellsPRMT1 272.6±6.5 23.3±0.6 8.3±0.2 1743±140 72PRMT2 122.3±3.2 51.8±1.5 3.7±0.1 157±14 78PRMT3 167.8±6.2 37.7±1.5 5.1±0.2 419±52 96PRMT4 261.0±3.0 24.2±0.2 7.9±0.1 1519±60 90PRMT5 641.2±33.5 10.0±0.6 19.5±1.1 23389±3729 100PRMT6 100.1±0.6 63.2±0.5 3.0±0.1 85±2 72PRMT7 116.8±2.5 54.2±1.3 3.6±0.1 137±10 115EGFP 70.3±0.9 90.2±1.1 2.1±0.1 29±1 30

Autocorrelation curves shown in Fig. 7 were fitted with the Zeiss FCS software module to determine the diffusion time t of individual PRMTs. All values aregiven as mean ± s.e.m. Using t values from untreated cells, the diffusion coefficients D, hydrodynamic radii rH and approximate mass of the molecules werecalculated as described in the Materials and Methods. The monomeric molecular mass of each PRMT is given for reference. Note that PRMT6 and PRMT7 (andthe control protein EGFP) have diffusion characteristics of monomeric proteins, PRMT2 behaves like a dimer, PRMT3 like a tetramer, and PRMT1 and PRMT4are present in large complexes. PRMT5 appears to aggregate or associate with very large structures.

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photobleaching experiments revealed that PRMT3 is the only PRMT

with almost identical diffusion coefficients in the nucleus and the

cytoplasm. This is unusual for a protein, because the viscosity of

the nucleus and the cytoplasm differs by a factor of two to three

(Wachsmuth et al., 2000), which is usually reflected in the diffusion

coefficient of a protein present in both compartments (see for

example, the mobility of GFP or PRMT7, Table 1A). The

comparatively slow diffusion of PRMT3 in the cytoplasm apparently

arises from interactions of the enzyme with substrates that are in

large complexes, presumably with rpS2 in ribosomes (Choi et al.,

2008; Swiercz et al., 2007). This interaction is not significantly

affected when methylation is inhibited, suggesting that methylation

of rpS2 (or other substrates) is not essential for binding. However,

the nuclear fraction of PRMT3 gains an immobile fraction under

Adox treatment, which is similar to PRMT1, but not as pronounced

(Fig. 3) (Herrmann et al., 2005). This suggests that nuclear PRMT3

interacts with entities of low mobility, possibly complexes

containing pre-mRNA and poly(A)-binding protein or nuclear

substructures, such as chromatin. Complementary in vitro

experiments demonstrated that human PRMT3 behaves like a

monomer or a small oligomer in glycerol gradients, in agreement

with gel filtration results reported by Tang and colleagues (Tang et

al., 1998). Quantitative analysis by fluorescence correlation

spectroscopy revealed diffusion characteristics compatible with

PRMT3 forming a tetramer in solution. Alternatively, PRMT3 could

be in a stable complex with another protein that affects its diffusion.

In this context, it is interesting to note that neither our gradient

centrifugations nor the fluorescence correlation spectroscopy

measurements revealed the presence of human PRMT3 bound to

ribosomal subunits, as previously described for PRMT3 from fission

yeast (Bachand and Silver, 2004) and mouse (Swiercz et al., 2007).

We obtained the same results for the endogenous and for the stably

expressed GFP fusion protein, suggesting that the interaction of

PRMT3 with ribosomes might be weaker in human cells.

Interestingly, while we were preparing this paper, Choi and

colleagues (Choi et al., 2008) published identical results for

PRMT3, also from human HEK293 cells. Thus, the binding of

human PRMT3 might be strong enough to be responsible for the

slow mobility of PRMT3 in the cytoplasm of HEK293 cells (see

above), but might dissociate when cells are disrupted.

PRMT7 is predominantly cytoplasmic in the HEK293 cells

investigated here, and is a very rapidly diffusing protein in vivo,

similar to PRMT3. Its diffusion characteristics in live cells are

typical for a protein with no or very few interactions with substrate

proteins, and they are not altered when methylation is inhibited.

Fluorescence correlation spectroscopy shows that the protein is

monomeric in solution. However, because PRMT7 has two

methyltransferase domains (see Fig. 1), the monomeric protein

might mimic oligomerization and behave as an intramolecular dimer.

This is in line with data from Miranda and colleagues who

demonstrated that both methyltransferase domains are required for

functionality (Miranda et al., 2004). In our experiments, we did not

detect enzymatic activity of PRMT7, suggesting that either the

specific activity of the expressed enzyme is too low (as is the case

for PRMT3) (Tang et al., 1998), or that the substrate is missing or

too dilute in the total cell extracts used for the activity assay. It is

possible that substrates of PRMT7 are also missing or too dilute in

the HEK293 cells, in contrast to PRMT1 and PRMT4 for example,

which have many substrates present at high concentrations, so that

photobleaching experiments measure only the free enzyme.

As stated above, our combined results provide novel and, for the

first time, quantitative data about the dynamics of PRMT family

members in human cells. They show that PRMTs have non-

overlapping properties, and are therefore probably involved in

different cellular processes. The data presented here provide a firm

ground for future experiments, which are certainly needed to better

understand the physiological roles of arginine methylation and to

find potential links to consequences of its misregulation.

Materials and MethodsCell culture and transfectionHuman embryonic kidney cells (HEK293), human osteosarcoma cells (U2OS), or

human cervix carcinoma cells (HeLa) were cultivated on plastic dishes in DMEM

with 10% fetal calf serum in a humidified atmosphere containing 5% CO2, and were

split 1:5 every second day. Human breast carcinoma cells (MCF-7) were cultured

identically, but in medium containing additional insulin at a final concentration of

10 μg/ml. Cells were transfected using polyethylenimine (Boussif et al., 1995) with

expression vectors encoding EGFP fusions of PRMT1 to PRMT8, respectively; stably

expressing cell lines were created by selection with G418 for 4-6 weeks and, after

selection, expressing cells were obtained by fluorescence-activated cell sorting of

GFP-positive cells. For FRAP analysis and microscopy, we used cell populations

rather than single cell clones, to rule out clonal variation or other potential problems

inherent in the work with single cell clones. The generation of the GFP-PRMT1 to

GFP-PRMT6 constructs (Frankel et al., 2002) and GFP-PRMT8 construct (Lee et

al., 2005a) has been described previously. Human cDNA was used to amplify PRMT7

by PCR. The PCR product was digested with BglII and EcoRI, and subcloned into

pEGFP-C1.

For inhibition of methylation and for preparation of hypomethylated cell extracts,

cell culture medium was supplemented with 15 μM of periodate-oxidized adenosine

(adenosine-2�-3�-dialdehyde, Sigma A7154), and cells were cultured for additional

48 hours before they were analyzed or harvested. Conditions of Adox treatment had

been empirically determined and thoroughly controlled as described (Herrmann et

al., 2005).

Live-cell microscopyFor live-cell microscopy, cells were split onto 35 mm culture dishes with a glass

bottom (Mattek) and analyzed 2 days after splitting either untreated or treated with

Adox as described above. For the analysis, cells were placed on the heated stage of

a Zeiss Meta 510 confocal laser-scanning microscope and images were acquired from

typical cells using a �63 oil-immersion lens and 488 nm excitation at 1% laser

intensity.

Fluorescence recovery after photobleaching (FRAP)Photobleaching was performed as described previously (Herrmann et al., 2005;

Mearini and Fackelmayer, 2006). Approximately 20 cells were analyzed for each

PRMT, and mean pixel intensities in the bleached region of interest (ROI) and a non-

bleached reference region were determined with the freeware image analysis software

ImageJ by Wayne Rasband (http://rsb.info.nih.gov/ij/). Raw recovery data were

imported into Microsoft Excel for normalization with pixel intensities from the

reference region and the initial fluorescence intensity directly after bleaching. Non-

linear regression was performed with Prism4 (Graphpad) to determine the mobile

fraction (Mf) and halftime of recovery (t/2). All PRMT recovery curves could

consistently be fitted very well with the Weibull function:

F(t) = Fmax*[1 – exp(–K*t^s)] ,

where F(t) represents the fluorescence at time t, Fmax represents the asymptote of the

ROI fluorescence recovery curve (equal to Mf), K represents the rate constant of

recovery, and s represents the Weibull ‘shape parameter’. This parameter describes

a change of the exponential recovery rate with time; for a constant recovery rate, sequals 1 and the Weibull function converts to the one-phase exponential association

function:

F(t) = Fmax*[1 – exp(–K*t)] .

For our experimental setup, a value of 0.74 for s yielded a significantly better

curve fitting than the one-phase exponential association function (see supplementary

material Fig. S1). Diffusion coefficients D (expressed as μm2/second) were then

calculated from t/2 using:

D = ω2 / (4*τ) ,

where τ represents t/2 in seconds, and ω represents the height of the bleaching region

in μm (Axelrod et al., 1976).

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Fluorescence correlation spectroscopyCell extracts were prepared by swelling cells stably expressing EGFP fusion proteinsin hypotonic buffer (20 mM Tris-HCl, pH 7.4) for 5 minutes, and homogenizationin a Dounce homogenizer (Wachsmuth et al., 2000). The extract was cleared bycentrifugation for 5 minutes at 6000 g, adjusted to isotonic conditions (20 mM Tris-HCl, pH 7.4, 130 mM NaCl) and diluted in the same buffer to 1-5 molecules perconfocal volume of measurement.

The FCS measurements were performed on the Zeiss LSM 510/ConfoCor 2combination. EGFP was excited with the 488 nm line from an argon-ion laser (1%laser intensity) and focused into the sample with a water immersion C-Apochromat�40 lens objective (Zeiss). The excitation and emission light were separated by adichroic beam splitter (HFT 488/633). The fluorescence was detected by an avalanchephotodiode after being filtered through a longpass filter >505 nm. The pinhole wasset to 50 μm. Between three and six independent measurements with 100autocorrelation traces (10 second measuring time) were performed for eachinvestigated protein. The data were analyzed using the Zeiss ConfoCor2 software.The autocorrelation traces were fitted with a model describing Brownian motion ofone or two distinct species in three dimensions (autocorrelation function). This curvefit resulted in values for τd, which represent the characteristic diffusion time of themolecules, and allow calculation of the translational diffusion coefficient D by usingthe equation:

D = ω2 / (4*τd) ,

where ω is the equatorial radius of the confocal measuring volume and is obtainedfrom calibration measurements using Rhodamine-6-green (Rh6G). In our experimentalsetup, ω was determined to be 159 nm.

Approximate values for the hydrodynamic radii were calculated, assuming aglobular shape of the molecules, using the Stokes-Einstein relation:

rH = k*T / (D*6π*η) ,

where rH represents the hydrodynamic radius of the molecule, k represents theBoltzmann constant, T represents the temperature (310 K), and η represents theviscosity of the solution which was experimentally determined from measurementswith EGFP (1.18 mPa seconds). The hydrodynamic radius allows estimating themolecular mass of the protein by using equation:

m = 4π*ρ*NA*rH3 ,

where ρ represents the mean density of the protein (set to 1.2 g/cm3) and NA representsAvogadro’s number.

Western blottingWestern blotting was performed as described previously (Herrmann et al., 2004) usingcommercially available antibodies against GFP (Roche, Applied Science), hnRNP-C (ab10294, Abcam) or PRMT1 to PRMT7 (07-404, 07-256, 07-405; 07-639, Upstate;ab3667, Abcam; IMG-496, IMG-506, Imgenex). For detection, HRP-conjugatedsecondary antibodies (Sigma) and ECL chemiluminescent reagent (AmershamBiosciences) were used.

In vitro methylation assaySemiconfluent to confluent cells were harvested by scraping off the culture disheswith a rubber policeman after two washes with PBS, and collected by centrifugation(200 g, 5 minutes). Extract was prepared by resuspending cells in lysis buffer [1.5�PBS, 1% Triton X-100 and protease inhibitor cocktail Complete, EDTA-free (Roche)],incubating on ice for 5 minutes, and clearing by centrifugation for 15 minutes in anEppendorf microcentrifuge at full speed. Extract from 1-5�106 cells wassupplemented with 5 μg antibodies against GFP (Roche, Applied Science), incubatedfor 2 hours at 4°C, and immune complexes were collected by incubation with protein-G-Sepharose for an additional 1 hour. Immune complexes were washed thoroughlysix times and the activity of precipitated PRMTs was measured by radioactivemethylation assays exactly as described earlier (Herrmann et al., 2004). Briefly,immunoprecipitated PRMT (on 20 μl Protein-G-Sepharose beads) were combinedwith heat-inactivated extract from hypomethylated cells and 5 μCi S-adenosyl-L-[methyl-3H]methionine (Amersham Biosciences TRK865, specific activity 2.96TBq/mmol), and reactions were incubated for 2 hours at 37°C. The reaction mixturewas then resolved by SDS-PAGE and radioactively labeled proteins were visualizedby fluorography with enHance reagent (NEN Dupont).

Glycerol gradient centrifugationNative total cell extract was prepared as described for in vitro methylation assay, andlayered over a 10-30% glycerol gradient (in lysis buffer) in a Beckman SW41 tube.The gradient was centrifuged at 4°C for 20 hours at 30,000 r.p.m. and was fractionatedin 0.8 ml aliquots from the top. Aliquots of each fraction were resolved by SDS page,and proteins were detected by western blotting.

We wish to thank Arne Düsedau from the Heinrich-Pette-Institute,Hamburg, Germany, for expert help with cell sorting and MagdalenaHyzy, Gdansk, for pioneering work with PRMT2. This work was

supported by grant Fa376/3-1 of the Deutsche Forschungsgemeinschaft(DFG) to F.O.F. and a short-term fellowship from the Scheringfoundation to Magdalena Hyzy.

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