Identification of a second DNA binding site in human DNA methyltransferase 3A by substrate inhibition and domain deletion

Archives of Biochemistry and Biophysics 498 (2010) 13–22

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Original paper

Identification of a second DNA binding site in human DNA methyltransferase 3A by substrate inhibition and domain deletion Matthew M. Purdy a,1, Celeste Holz-Schietinger b, Norbert O. Reich a,b,* a b

Department of Chemistry and Biochemistry, University of California, Santa Barbara, CA 93106-9510, USA Program in Biomolecular Science and Engineering, University of California, Santa Barbara, CA 93106-9510, USA

a r t i c l e

i n f o

Article history: Received 16 December 2009 and in revised form 6 March 2010 Available online 17 March 2010 Keywords: DNA methylation Protein–DNA interaction Allosteric binding Enzyme kinetics Non-specific DNA binding

a b s t r a c t The human DNA methyltransferase 3A (DNMT3A) is essential for establishing DNA methylation patterns. Knowing the key factors involved in the regulation of mammalian DNA methylation is critical to furthering understanding of embryonic development and designing therapeutic approaches targeting epigenetic mechanisms. We observe substrate inhibition for the full length DNMT3A but not for its isolated catalytic domain, demonstrating that DNMT3A has a second binding site for DNA. Deletion of recognized domains of DNMT3A reveals that the conserved PWWP domain is necessary for substrate inhibition and forms at least part of the allosteric DNA binding site. The PWWP domain is demonstrated here to bind DNA in a cooperative manner with lM affinity. No clear sequence preference was observed, similar to previous observations with the isolated PWWP domain of Dnmt3b but with one order of magnitude weaker affinity. Potential roles for a low affinity, low specificity second DNA binding site are discussed. Published by Elsevier Inc.

Introduction DNA methylation plays key roles in regulating gene expression and maintaining genomic stability [1,2]. Establishing patterns of DNA methylation to silence imprinted genes and certain repetitive elements is necessary for embryonic development, and abnormal DNA methylation patterns are found in many types of cancer and developmental disorders. More recently, changes in DNA methylation have been linked to memory formation [3–5] along with infection by HIV and other viruses [6–9]. DNA methylation in mammals is established and maintained after replication by the DNA methyltransferase (DNMT)2 family of enzymes, which methylate the cytosine in CG target sites with only modest preferences for the sequence of bases flanking this dinucleotide. Given that methylation occurs in a highly tissue-specific and developmental stage-specific manner, understanding how these enzymes are regulated and targeted to specific genomic regions is of great interest.

* Corresponding author. Address: Department of Chemistry and Biochemistry, University of California, Santa Barbara, CA 93106-9510, USA. Fax: +1 805 893 4120. E-mail address: [email protected] (N.O. Reich). 1 Present address: Affymetrix, Inc., 3450 Central Expressway, Santa Clara, CA 95051, USA. 2 Abbreviations used: DNMT3A, human DNA methyltransferase 3A (full length); Dnmt3a, murine DNA methyltransferase 3a; Dnmt1, murine DNA methyltransferase 1; AdoMet, S-adenosyl-L-methionine; PCR, polymerase chain reaction; PMSF, phenylmethanesulfonyl fluoride; EDTA, ethylenediaminetetraacetic acid; DTT, DL-dithiothreitol; BME, 2-mercaptoethanol; BSA, bovine serum albumin; SDS, sodium dodecylsulfate; PAGE, polyacrylamide gel electrophoresis; EMSA, electrophoretic mobility shift assay; SEC, size exclusion chromatography. 0003-9861/$ - see front matter Published by Elsevier Inc. doi:10.1016/j.abb.2010.03.007

The murine Dnmt1 isoform shows a preference for methylation of hemimethylated DNA (where one strand of the target CG site is already methylated) [10–13], which is consistent with a major role in copying the methylation pattern after replication. The DNMT3A isoform and the closely related DNMT3B are considered to play the main roles in establishing new methylation patterns during embryonic development [1,2]. These enzymes lack the preference for hemimethylated sequences and are expressed in various splice forms at different developmental stages. The division of maintenance and de novo methylation activities between DNMT1 and the DNMT3 enzymes is not believed to be absolute [2,14]. The active site residues of the C-terminal catalytic domain, which is responsible for methyltransferase activity, are conserved in DNMT1 and the DNMT3 enzymes [14,15]. The N-terminal regulatory regions of DNMT1 and the DNMT3 enzymes differ significantly. In addition to the catalytic domain, DNMT3A and DMNT3B contain nuclear localization signals [16], a PWWP domain, a CXXC/PHD domain, and a non-conserved N-terminal region (Fig. 1) [14]. The PWWP domain family, named for its conserved tryptophan-tryptophan containing motif, is found in several proteins involved in transcriptional regulation and chromatin organization [17]. In the noncatalytic DNMT3L, the zinc-binding CXXC/PHD domain is important for the recognition of histone H3 [18]. In the DNMT3 enzymes, the Nterminal region, PWWP domain, CXXC/PHD domain, and catalytic domain have all been implicated in mediating protein–protein interactions, with some of these interactions suggested to be involved in enzyme targeting [7,19–23]. The PWWP domain of Dnmt3b, along with that of the hepatoma-derived growth factor [24], binds DNA non-specifically [16,25]. This domain was shown to be necessary

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PWWP PHD/ Catalytic CXXC

Full length (1-912)

1

DNMT3A2 (Δ1-223)

912 224

ΔPWWP (Δ1-459)

912 460

Catalytic domain (Δ1-611) PWWP domain (279-434)

912 612

279

912

434

Fig. 1. Truncations of DNMT3A used in this study. Schematic of DNMT3A with PWWP domain in black solid, PHD/CXXC domain in black stripes, and catalytic domain in gray.

for localizing Dnmt3a and Dnmt3b to chromatin, but the role of the PWWP domain’s DNA binding ability in this targeting is debatable [16,26]. Substrate inhibition studies showed that Dnmt1 contains a second DNA binding site outside of its active site [10,27–32]. The discovery [29] and mechanistic characterization [30,31] of a very potent single stranded DNA inhibitor binding at this as yet unidentified allosteric site of Dnmt1 has led to the proposal that nucleic acid binding may regulate the methyltransferase activity. This is of potential interest because of the recent realizations of the importance of non-coding RNA molecules in regulating gene expression [33–35], DNMT’s as drug targets [29,36,37], and advantages of targeting inhibitors to allosteric sites [38–40]. This work addresses the ability of DNMT3A’s PWWP domain to bind a second, non-substrate DNA molecule and the functional consequences of occupancy of this binding site. Materials and methods Materials

Table 1 Oligonucleotide sequences used in this study.a Name

Sequence

GCbox2

50 -GGGAATTCAAGGGGCGGGGCAATGTTAGGG-30 30 -CCCTTAAGTTCCCCGCCCCGTTACAATCCC-50 50 -GGGAATTCAAGGGGMGGGGCAATGTTAGGG-30 30 -CCCTTAAGTTCCCCGCCCCGTTACAATCCC-50

GCbox2_hmB

50 -GGGAATTCAAGGGGCGGGGCAATGTTAGGG-30 30 -CCCTTAAGTTCCCCGMCCCGTTACAATCCC-50

GCbox2_fm

50 -GGGAATTCAAGGGGMGGGGCAATGTTAGGG-30 30 -CCCTTAAGTTCCCCGMCCCGTTACAATCCC-50

EF1a

50 -GGATGAAGGTGGCGCGGGGTAAACTGAAGG-30 30 -CCTACTTCCACCGCGCCCCATTTGACTTCC-50

Abox

50 -GGGAATTCATGGCGCAGTGGGTGGATCCAG-30 30 -CCCTTAAGTACCGCGTCACCCACCTAGGTC-50

CDH

50 -GGGAATTCAACCCCACTGCCCCTGTCCG CCCCGACTTAATGTTAGGG-30 30 -CCCTTAAGTTGGGGTGACGGGGACAG GCGGGGCTGAATTACAATCCC-50

a b

DNMT3A and truncation construction The full length DNMT3A was PCR amplified from a cDNA clone (kindly provided by En Li and Taiping Chen, Novartis Institutes for Biomedical Research) using the primers 50 -CCGTCGCCCATATGCCC GCCATGC-30 and 50 -GTTTGCCCCGCGGCCGCTTACACACACG-30 to append NdeI and NotI restriction sites as underlined. The PCR product was digested with NdeI and NotI then ligated into a similarly digested pET28a vector. The final 965 bp (from an internal KpnI restriction site to the stop codon) of the native coding sequence for DNMT3A were then replaced by a synthetic gene which had been codon optimized for expression in Escherichia coli (a generous gift from Frédéric Chédin, University of California at Davis, unpublished results) to produce the vector pET28-DNMT3ACopt, which codes for expression of the 912 amino acids of DNMT3A preceded by the Nterminal tag sequence MGSSHHHHHHSSGLVPRGSH. Constructs for expressing the DNMT3A truncations shown in Fig. 1 were created by amplifying selected regions out of pET28-DNMT3ACopt and subcloning into the NdeI and NotI restriction sites of pET28a to produce truncations coding for the noted amino acids preceded by the same N-terminal tag sequence. The rationale behind these truncation divisions is described in the Supplementary Materials section. Protein expression and purification

The DNA oligonucleotides shown in Table 1 were obtained from Midland Certified Reagent Company (Midland, TX) and purified by

GCbox2_hmA

reverse phase HPLC as described [41], except the strands of the EF1a duplex were purchased from Integrated DNA Technologies (Coralville, IA) and not HPLC purified. Duplexes were prepared by annealing the complementary strands in a pH 8.0 buffer containing 10 mM Tris–Cl, 1 mM EDTA, and 50 mM NaCl. Unlabeled S-adenosyl-L-methionine (AdoMet) was a product of Sigma–Aldrich; S-[methyl-3H]adenosyl-L-methionine and polydeoxy(inosinatecytidylate) acid, sodium salt (poly dIdC) were purchased from GE Lifesciences. Stock solutions of poly dIdC were quantified using the 260 nm extinction coefficient of 6.9  103 M1 cm1 provided on the manufacturer’s specification sheet and heated at 45 °C for 5 min before use in the assays.

CL2

50 -CAACAACTTCTTCTTCTTCTTCTTCTTCTTCTTCAACAAC-30 30 -GTTGTTGAAGAAGAAGAAGAAGAAGAAGAAGAAGTTGTTG-50

GCbox2topb

50 -GGGAATTCAAGGGGCGGGGCAATGTTAGGG-30

‘‘M” denotes 5-methyldeoxycytosine; CG target sites underlined. Single stranded oligonucleotide. All others are duplexes.

Proteins were expressed in E. coli strain Rosetta2(DE3)pLysS (from Novagen), except the catalytic domain was expressed in E. coli strain ER2566 (from New England Biolabs) that had been previously transformed with the pLysS plasmid (from Novagen). Shaker flask cultures were grown in 2 YT media at 37 °C to an OD600 nm of 0.7. The temperature was lowered to 28 °C and expression was induced by addition of 1 mM isopropyl-b-D-thiogalactopyranoside (Fisher Scientific). After shaking for 4 h at 28 °C, cells were harvested by centrifugation, washed with 20 mM potassium phosphate, 50 mM NaCl, pH 7.5 buffer, and frozen at 80 °C. For full length DNMT3A purification, cells were lysed by freezing and thawing followed by sonication to shear nucleic acids. The protein was purified by chromatography on BioRex-70 (BioRad), HiTrap Nickel-Sepharose (GE Lifesciences), and Superdex-200 prep grade (GE Lifesciences) resins. The Nickel-Sepharose column involved a lengthy wash with a buffer containing 0.1% Triton X-100 followed by a brief wash without this detergent before elution. Further details are given in the Supplemental Methods section. The protein was then dialyzed into 50 mM Tris–Cl, 200 mM NaCl, 1 mM EDTA, 20% (v/v) glycerol, pH 7.2 (when measured at room temperature) buffer with 0.5 mM DTT and stored at 80 °C. The DNMT3A truncations were purified by modifications of the above methods summarized here and described in greater detail in the Supplementary Materials section. The DNMT3A2 form and DPWWP truncation were purified first with a Nickel-Sepharose column and then using a BioRex-70 column. For the catalytic domain, the lysate was passed through a DEAE column then purified using a Nickel-Sepharose column, omitting Triton X-100 from the lengthy

M.M. Purdy et al. / Archives of Biochemistry and Biophysics 498 (2010) 13–22

wash step. The eluant was further purified on a BioRex-70 column as above. The PWWP domain was chromatographed on Nickel-Sepharose then further purified on a HiTrap SP Sepharose HP column (GE Lifescience). For all four DNMT3A truncations, the size exclusion chromatography (SEC) step was omitted. Protein purities of 80, 90, 80, 95, and 95% for (in order) full length DNMT3A (Fig. 2), DNMT3A2, the DPWWP truncation, the catalytic domain, and the PWWP domain were estimated by densitometry of Coomassie stained SDS–PAGE gels. The total protein concentrations in the full length DNMT3A and PWWP domain preparations were determined by Bradford assays using bovine serum albumin as a standard. These results were within 20% of the concentrations determined using 280 nm extinction coefficients calculated by the method of Gill and von Hippel [42] as implemented by the website http://www.scripps.edu/_cdputnam/ protcalc.html. The total concentration of full length DNMT3A was calculated by multiplying the total protein concentration by the fraction of purity. The concentrations of the DNMT3A2, DPWWP, and catalytic domain truncations were determined by 280 nm absorbance using calculated extinction coefficients.

inhibition constant, and b is the rate of turnover for the ESS complex divided by that for the ES complex. This equation was derived from the rapid equilibrium equation for non-competitive partial inhibition [45] by substituting the substrate concentration for the inhibitor concentration. Here Ks is used rather than Km to acknowledge the rapid equilibrium assumption. For the DNMT3A2 form with poly dIdC as a substrate, the data were fitted to a standard uncompetitive substrate inhibition equation (Eq. (2)) [46].

v¼

v¼



S Ks



2

þ KbS s K

i 2

1 þ KSs þ KSi þ K sS K i

ð1Þ

where m is the initial rate, S is the concentration of the DNA substrate, Ks is the substrate dissociation constant, Ki is the substrate

ð2Þ

i

v¼

V max

V max S   K m þ S 1 þ KS

Kinetic measurements of the full length DNMT3A with the GCbox2 duplex DNA substrate were employed to determine f , the fraction of active enzyme. The concentration of substrate was varied at concentrations close to that of the enzyme. The parameters kcat, Ks, and f for the full length DNMT3A with GCbox2 as a substrate were obtained by globally fitting m as a function of both S and E, the concentration full length DNMT3A (sum of both active and inactive, determined by the Bradford assay), to Eq. (3) using SigmaPlot.

Methylation assays Reactions were carried out at 37 °C in a buffer of 50 mM Tris–Cl and 1 mM EDTA at pH 7.8 (measured at room temperature) with 1 mM DTT, 0.2 mg/mL BSA, 25 mM NaCl, 2 lM AdoMet (a 6 Ci/ mmol mix of unlabeled and 3H methyl labeled). Reactions were initiated by addition of substrate DNA, then quenched after 1 h by addition of 500 lM unlabeled AdoMet and digestion with 50 lg/ mL proteinase K at 37 °C for 10 min (similar to Suetake et al. [43]). Samples were spotted onto Whatman DE81 filters then washed, dried, and counted as described previously [44], but omitting the 100% ethanol wash step. For the full length DNMT3A with poly dIdC as a substrate, the data were fitted to the partial substrate inhibition model shown in Eq. (1):

15

 qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi kcat f  E þ K s þ S  ðf  E þ K s þ SÞ2  4f  E  S 2

ð3Þ

In this equation S and E are the total concentrations (sum of both free and bound). The spirit of this method is similar to Dixon’s method to determine enzyme concentrations by quantifying the deviation from the Michaelis–Menten equation when the enzyme concentration is near that of the substrate [45,47]. Rather than employing Dixon’s graphical method, the rapid equilibrium assumption was made and the data were fitted to a quadratic equation for rapid equilibrium kinetics at high enzyme concentrations [45] in a similar manner as employed in determining the enzyme concentration in tight binding inhibition studies [48]. Rather than obtaining data from only one enzyme dilution, Eq. (3) was modified from a quadratic equation for rapid equilibrium kinetics at high enzyme concentrations [45] by substituting the term fE in for the enzyme concentration. This differentiates active from inactive enzyme, allowing global fitting at multiple enzyme concentrations, rather than fitting at each enzyme concentration separately followed by averaging. Substrate concentrations were chosen to be below the onset of substrate inhibition.

Fig. 2. Expression and purification of full length human DNMT3A and its truncations. (a) His6 tagged full length DNMT3A expressed in E. coli was purified by cation exchange, Ni-affinity, and size exclusion chromatographies. Lanes of an 8% acrylamide SDS–PAGE gel are: 1, crude lysate; 2, after BioRex-70 column; 3, after Nickel-Sepharose column; 4, after Superdex-200 column and dialysis; M, protein molecular weight markers. (b) Purified proteins: 1, catalytic domain (D1–611); 2, DNMT3A2 (D1–224). (c) Purified DPWWP (D1–459).

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DNA binding measurements Duplex oligonucleotide binding by the PWWP domain was measured by an electrophoretic mobility shift assay (EMSA) similar to that previously described [49]. Duplex DNA oligonucleotides GCbox2 (1.5 nM) or CDH (1.2 nM), with one strand 32P end labeled, were incubated with 0–10 lM PWWP domain for 20 min at room temperature in a buffer containing 50 mM Tris–Cl, 1 mM EDTA, pH 7.8 with 25 mM NaCl, 1 mM EDTA, 1 mM DTT, 0.2 mg/mL BSA, and 7% (w/v) glycerol. Samples were separated by non-denaturing PAGE (10% acrylamide in 1 TBE) run at room temperature, then imaged using phosphor storage screens and a Storm 840 phosphorimager (both from GE Lifesciences). The bands were quantified by densitometry using ImageJ [50] and fitted to the Hill equation using SigmaPlot. For a rapid comparison of the PWWP domain’s ability to bind various duplex and single stranded oligonucleotides (Fig. 8), 100 nM duplex or 300 nM single stranded oligonucleotides were incubated with 0, 0.5, or 5 lM PWWP domain and separated on 8 or 10% acrylamide gels in 0.5 TBE. These gels were then stained with Sybr Gold (Invitrogen) and imaged on a Storm 840 phosphorimager.

Results Protein purification and enzyme activity The full length human DNMT3A was expressed in E. coli with an N-terminal His6 tag and purified to approximately 80% homogeneity (Fig. 2). A portion of the DNMT3A coding sequence utilizes a synthetic gene which had been codon optimized for expression in E. coli (a generous gift from Frédéric Chédin, University of California at Davis, unpublished results). The protein eluted just after the void volume of the size exclusion chromatography (SEC) column though this 912 amino acid, 102 kDa protein should fit well into the Superdex-200 resin’s 10–600 kDa fractionation range. This suggests that full length DNMT3A forms high molecular weight aggregates as observed for DNMT3A2 [51,52]. DNMT3A eluted in the void volume of the SEC column even in the presence of 2 M NaCl as indicated previously for DNMT3A2 [51] and in the presence of 0.1% Triton X-100. Western blotting with antibodies to a His6 tag and to residues 469–483 of DNMT3A showed that the major contaminants (Fig. 2) are N-terminal fragments of DNMT3A (data not shown). These catalytic domain lacking N-terminal fragments are not expected to contribute to catalytic turnover, but co-elution with DNMT3A from the SEC column suggests that the fragments may be aggregated with the full length enzyme. We cannot rule out the possibility that aggregation with these fragments diminishes the activity of the full length enzyme, though it seems unlikely that aggregation with fragments would diminish the activity any more than aggregation with other full length molecules. Several truncations of DNMT3A (Fig. 1) were also expressed and purified (Fig. 2). Washing the protein-bound Nickel-Sepharose columns with the detergent Triton X-100 was found to be necessary for avoiding precipitation of the DNMT3A2 and DPWWP forms during later purification stages. Precipitation of the catalytic domain (which was not subjected to the detergent wash) was noted when 2–4 lM stock solutions were warmed to room temperature. However, under assay conditions with poly dIdC as a substrate, the catalytic domain showed a linear progress curve for at least 90 min (data not shown), suggesting that complexation with co-factor or substrate stabilizes this truncation of DNMT3A. The percentage of active DNMT3A was estimated kinetically using a 30-mer duplex DNA substrate (Fig. 3). This quantifies the enzyme concentration by examining deviations from hyperbolic

Fig. 3. Determination of fraction of active enzyme in the DNMT3A preparation. Initial rate measurements were determined at 100 nM (), 200 nM (+), and 300 nM (s) DNMT3A protein concentrations (as determined by a Bradford assay) at varying concentrations of the GCbox2 duplex DNA substrate under the conditions described in Fig. 4. Data points are averages of duplicate measurements. Data for rate as a function of both DNA and protein concentrations were simultaneously fit to the quadratic equation (Eq. (3)) resulting in values of 0.43 (±0.03) h1, 2.6 (±1.4) nM, and 0.18 (±0.01) for kcat, Ks(GCbox2), and f (the fraction of active enzyme), respectively. For illustration purposes, the data were plotted in two dimensions, and curves were simulated for each protein concentration using the above values from the global fit.

binding when the enzyme concentration is close to that of the substrate. Here substrate concentrations close to the enzyme concentration were varied and the process repeated at several different enzyme concentrations. Qualitatively, one can see in Fig. 3 that the measured protein concentrations cannot be a reasonable estimate of the active enzyme concentrations. For the curve marked by open circles, if the 300 nM protein concentration employed there were all active enzyme, 150 nM would be the lowest substrate concentration at which the half-maximal rate could occur. However, the curve is nearly saturated at 150 nM substrate. To quantify this observation, the data were fitted to a quadratic kinetic expression (Eq. (3)). The rapid equilibrium assumption required for use of this equation may be reasonable given the very slow rate of enzymatic turnover observed with the DNMT3 enzymes. This fitting procedure estimates that only 18% of the full length DNMT3A is catalytically active. To facilitate comparisons between this full length DNMT3A preparation, the DNMT3A truncations (for which the active enzyme fraction was not measured), and DNMT3 forms from previous studies [43,51,53–56], the DNMT3A concentrations used here are given in terms of protein concentration rather than active enzyme (except as noted). Mechanism of substrate inhibition by poly dIdC High poly dIdC concentrations inhibit full length DNMT3A (Fig. 4a). Substrate inhibition can occur by binding of a second DNA molecule outside of the enzyme active site if the resulting enzyme–DNA–DNA complex turns over more slowly than the enzyme–DNA complex. It could also result from formation of an enzyme–DNA–AdoHcy complex.3 The data in Fig. 4a show partial substrate inhibition, where the reaction rate is not forced to zero at an infinite substrate concentration. This is consistent with the former mode of inhibition but also consistent with the latter if there is a random order of product release. Prior inhibition studies on Dnmt3a support an ordered mechanism where AdoMet binds second and AdoHcy is released first [53]. This would not predict formation of 3 In this and the following paragraph, DNA refers to the unmethylated (substrate) form of the DNA as opposed to the methylated (product) form; AdoHcy is the product form of AdoMet.

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Fig. 4. Substrate inhibition of DNMT3A truncations by poly dIdC. Initial rates were measured with poly dIdC as a substrate at 37 °C with 2 lM AdoMet in a buffer containing 50 mM Tris–Cl, pH 7.5, 25 mM NaCl, 1 mM EDTA, 1 mM DTT, and 0.2 mg/mL BSA. (a) 30 nM full length DNMT3A with poly dIdC concentrations from 0.3 to 400 lM; (b) 40 nM DNMT3A2 (D1–224) with poly dIdC concentrations from 0.25 to 300 lM; (c) 50 nM DPWWP (D1–459) with poly dIdC concentrations from 0.25 to 300 lM; (d) 30 nM catalytic domain (D1–611) with poly dIdC concentrations from 0.25 to 300 lM. Full length DNMT3A and truncation concentrations are total protein concentrations based on a Bradford assay or 280 nm absorbance. Poly dIdC concentrations are reported in base pairs. Data points are the average of three replicate measurements for (a) and a combination of triplicate and duplicate measurements for (b), (c), and (d). The inset in (a) shows the low substrate concentrations on an expanded scale. Data were fitted to the partial substrate inhibition model give in Eq. (1) for (a), a standard uncompetitive substrate inhibition model (Eq. (2)) for (b), or to a rectangular hyperbola for (c) and (d).

an enzyme–DNA–AdoHcy complex and supports the binding of a second DNA molecule as the cause of substrate inhibition. The model where inhibitory E–DNA complexes form in an ordered mechanism and AdoMet must bind to the enzyme first can be ruled out. Substrate inhibition by the DNA would disappear upon saturation with AdoMet. Also, this model would not predict partial inhibition. Both of these features are shown in the steady state rate equation given as Eq. (1) of Praest et al. [57]. Domain deletion was employed to gain more direct evidence for the existence of a second DNA binding site in DNMT3A. Fig. 4d shows that the isolated catalytic domain of DNMT3A (D1–611) lacks substrate inhibition by poly dIdC, even at a concentration of more than 3000 times its Km. This strongly suggests that the substrate inhibition observed in the full length enzyme results from a second DNA molecule occupying a second binding site located in the first 611 residues of DNMT3A. We cannot entirely rule out formation of a putative Enzyme–DNA–AdoHcy complex as the cause of substrate inhibition, for it is possible that truncation of the protein could alter active site DNA binding in a putative inhibitory Enzyme–DNA–AdoHcy complex more than in the catalytic Enzyme–DNA–AdoMet complex. Location of the second DNA binding site Two more truncations of DNMT3A were employed to further locate the second DNA binding site. Removal of the first 223 amino acid residues creates the DNMT3A2 splice form but leaves both the PWWP and PHD/CXXC regions intact. This form retains the substrate inhibition at high poly dIdC concentrations (Fig. 4b). The DPWWP truncation removes the first 459 residues of the protein, but retains the PHD/CXXC domain. This truncation lacks substrate inhibition (Fig. 4c), suggesting that PWWP domaincontaining region of residues 224–459 is necessary in forming the second DNA binding site.

Kinetic parameters with poly dIdC as a substrate The data in Fig. 4a were fitted to a rapid equilibrium model for partial non-competitive substrate inhibition (Eq. (1)) to give catalytic parameters of the full length DNMT3A (Table 2). The parameter ‘‘b” from Eq. (1) represents the ESS complex turnover rate constant divided by kcat (the ES complex turnover rate constant). In this model, a b value of 0.45 indicates that binding a second DNA molecule inhibits DNMT3A turnover approximately 2-fold. Table 2 Kinetic parameters for full length DNMT3A and truncations with poly dIdC as the substrate.a Full length DNMT3A kcat (h1) Relative activityb Km (lM)c,d Ki (lM)c bf

1.36 ± 0.09 1 1.4 ± 0.2 100 ± 70 0.45 ± 0.09

DNMT3A2

DPWWP

Catalytic domain

0.40 ± 0.04 0.84 ± 0.27 400 ± 160 n.m.e

0.34 ± 0.02 0.95 ± 0.15 n.m.e n.m.e

0.48 ± 0.04 0.47 ± 0.13 n.m.e n.m.e

a Apparent steady-state kinetic parameters were determined at 2 lM AdoMet from data and fitting described in Fig. 4; enzyme concentrations determined by a Bradford assay or 280 nm absorbance; if the fraction of active enzyme determined from Fig. 3 were taken into account, the kcat value for the full length DNMT3A would be 7.5 (±0.5) h1. b For the truncations, activity relative to wild type [(kcat for truncation)/kcat for full length)] are reported rather than kcat values due to possible difference in the stabilities and fractions of active enzymes in the truncations. c Km, and Ki values are for poly dIdC; concentrations are in lM of base pairs. d For full length DNMT3A, Km is more accurately called Ks, the substrate dissociation constant, due to the rapid equilibrium assumption used in the model from which Eq. (1) is derived. e Not meaningful. f From Eq. (1); represents turnover rate with inhibitory substrate bound divided by kcat (turnover rate without inhibitory substrate bound). In this model, a b value of 0.45 indicates that binding a second DNA molecule inhibits DNMT3A turnover approximately 2-fold.

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For the DNMT3A2 form, the data in Fig. 4b could not distinguish between partial and full substrate inhibition models and were fitted to the more standard uncompetitive substrate inhibition equation, in which the reaction rate approaches zero at an infinite substrate concentration. The Michaelis–Menten kinetic parameters for the DPWWP truncation and catalytic domain of DNMT3A are also shown in Table 2. The kinetic parameters in Table 2 indicate that there is no great loss of catalytic function in any of the DNMT3A truncations tested. The kcat values are difficult to compare directly to the full length enzyme because of potential instability of the truncations. Also, the truncations may have different fractions of active enzyme than the full length. The value of Km(poly dIdC) determined for the catalytic domain is lower than that for the full length enzyme. The reason for this is currently unclear. Substrate inhibition by oligonucleotide duplexes Findings with poly dIdC were confirmed using duplex oligonucleotide substrates containing one or two CpG target sites per strand. Using the 30-mer duplex GCbox2 with one CpG site per strand, the full length DNMT3A and DNMT3A2 truncation undergo substrate inhibition, while the isolated catalytic domain does not (Fig. 5). For the full length DNMT3A, the rate peaks at approximately 80 nM duplex, while the total enzyme concentration is 200 nM. This agrees with the finding that only a small fraction of the enzyme is in an active form (Fig. 3). Several other DNA duplexes were tested, and all show substrate inhibition of DNMT3A over the 2–5 lM range of duplex concentrations (Fig. 6). The oligonucleotide substrate Km(DNA) values were too small to reliably gather the low substrate concentration data necessary for fitting to a substrate inhibition model. The hemimethylated duplex GCbox2_hmB is a very poor substrate relative to its unmethylated counterpart or even GCbox2_hmA, where the methyl group is moved to the other strand (see Table 1 for oligonucleotide sequences). The GCbox2_hmA presents a CCCGC site for methylation, fitting into the preference of C/T-NC-G-C/T for Dnmt3a determined previously [56]. The GCbox2_hmB duplex presents a GGCGG site, which conforms poorly to this flanking sequence preference. Also, the best oligonucleotide substrate tested here, the 30-mer EF1a from the elongation factor 1a promoter, contains two CpG sites per duplex conforming to this site preference pattern.

Fig. 6. Substrate inhibition of full length DNMT3A is observed with several oligonucleotide sequences. Rates were measured using the 30- or 47-mer duplex DNA substrates: EF1a (j), Abox (N), CDH (d), GCbox2_hmA (s), and GCbox2_hmB (+). Conditions as in Fig. 4 with 100–250 nM full length DNMT3A (as determined by a Bradford Assay).

DNA binding by PWWP domain Previously, the PWWP domain of murine Dnmt3b was shown to bind DNA with a modest affinity of 230 nM [25]. The human DNMT3A PWWP domain is 51% identical to its murine Dnmt3b counterpart, but several basic side chains from the proposed binding surface [25] are not shared by DNMT3A. DNA binding by the PWWP domain of DNMT3A (residues 279–434) was measured by an electrophoretic mobility shift assay (EMSA) with radiolabeled duplex DNA oligonucleotides (Fig. 7). Cooperative binding was observed and several possible reasons for this are described in the Discussion section. Fits to the Hill equation indicated KD values of 3.5 (±0.2) lM for the CDH 47-mer and 6.1 (±0.4) lM for the GCbox2 30-mer with Hill coefficients of 2.1 (±0.1) and 2.8 (±0.3), respectively. The reason for the incomplete free DNA band shifting is unclear. Constraining the fits to 100% maximal shifts resulted in only small increases in KD values with small decreases in Hill coefficients. The murine Dnmt3b PWWP domain was previously suggested to bind DNA non-specifically [16]. DNMT3A PWWP domain binding to several DNA duplex oligonucleotides was tested using a more rapid EMSA technique with detection using a stain for DNA. The effect of protein complexation may interfere with DNA staining, but focusing on the free DNA bands in Fig. 8 indicates that at least 50% is shifted upon addition of 5 lM PWWP domain for the four different duplex DNA sequences tested (GCbox2, CDH, EF1a, and CL2). The CL2 sequence does not contain any CpG recognition sites (Table 1). In addition, hemimethylated and doubly methylated versions of the GCbox2 substrate were also at least 50% bound at 5 lM PWWP domain. Hybrid duplexes with one strand of DNA hybridized to one strand of RNA were also bound by the DNMT3A PWWP domain in this assay (data not shown). Very little binding was observed for a single strand of the GCbox2 sequence, which lacks predicted secondary structure [58]. This indicates that, though DNA binding by the PWWP domain is fairly non-specific, it does not recognize every polyanion. There appears to be a preference for duplex structures. The PWWP domain of mismatch repair protein MSH6 shows a similar preference for binding duplex over single stranded DNA [59]. Discussion

Fig. 5. Substrate inhibition of DNMT3A truncations by oligonucleotide substrate. Rates measured using the 30-mer DNA duplex GCbox2 as a substrate with other conditions as in Fig. 4. Protein concentrations were: 200 nM full length DNMT3A (s); 300 nM DNMT3A2 (+); 300 nM catalytic domain (N). The DPWWP truncation was not tested with this substrate. Full length DNMT3A and truncation concentrations stated here are total protein concentrations.

Comparison with previous studies The full length human DNMT3A was expressed in E. coli, purified, and characterized in this work. A turnover rate constant of

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Fig. 7. Isolated PWWP domain of DNMT3A can bind DNA. Binding of the isolated PWWP domain of DNMT3A (residues 279–434) to duplex DNA oligonucleotides was monitored by an electrophoretic mobility shift assay. Samples containing 1 nM 32P labeled DNA were incubated with 0–10 lM PWWP domain at room temperature in a buffer containing 50 mM Tris–Cl, pH 7.8, 25 mM NaCl, 1 mM EDTA, 1 mM DTT, 0.2 mg/mL BSA, and 7% (w/v) glycerol then separated by non-denaturing PAGE. (A and B) Autoradiographs of the gels with increasing concentrations of PWWP domain from left to right with the 30-mer GCbox2 DNA in (A) and the 47-mer CDH DNA in (B). (C) Densitometry results from the two gels; data from GCbox2 (d) and CDH (s) were fitted to the Hill equation resulting in KD values of 3.5 (±0.2) lM for CDH and 6.1 (±0.4) lM for GCbox2 with Hill coefficients of 2.1 (±0.1) and 2.8 (±0.3), respectively.

Fig. 8. PWWP domain binds DNA non-specifically. The binding of duplex and single stranded DNA sequences to the isolated PWWP domain were monitored by an electrophoretic mobility shift assay. Bands were detected by Sybr Gold staining of the DNA. Other conditions as in Fig. 7. Oligonucleotide sequences described in Table 1. (a) Lanes 1–2: 100 nM GCbox2 duplex DNA; lanes 3–4: 100 nM GCbox2_fm fully methylated duplex DNA; lanes 5–7: 100 nM CL2 duplex DNA (no CG sites). (b) Lanes 1–3: 100 nM GCbox2_hmA hemimethylated duplex DNA; lanes 4–6: 100 nM CDH duplex DNA; lanes 7–9: 100 nM EF1a duplex DNA; lanes 10–12: 300 nM GCbox2top single stranded DNA.

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1.2 h1 was measured with poly dIdC as a substrate. The turnover rate constant is defined here as the maximal rate from Fig. 4a divided by the protein concentration. This value falls into the range reported for murine Dnmt3a of 1.8 and 1.07 h1 for proteins expressed in insect cells [43] and E. coli [55], respectively, and for the human DNMT3A2 splice variant of 0.52 h1 [51] expressed in E. coli. In addition, murine Dnmt3a expressed in mammalian cells had a turnover rate constant of >0.18 h1 using a plasmid DNA substrate [56] (estimated from initial rate in Fig. 5c of the reference). These comparisons suggest that the observation of the low fraction of active enzyme in the present preparation is not unique to the human DNMT3A or E. coli expression, although prior studies did not report the fraction of DNMT3A which was active. The self aggregation of DNMT3A may not merely be a function of over-expression in E. coli [51]. Crystallographic evidence combined with mutagenesis points to a functional oligomer [52,60]. This aggregation phenomenon may be related to the high percentage of inactive enzyme. Thus the large, heterogeneous aggregates observed in solution [51,52,60] may be inactive, and only a small percentage of DNMT3A may exist as small, active oligomers. Alternatively, all of the enzyme may exist as aggregates and only a small number of protomers in each aggregate may be active. In support of the former, inclusion of 0.1% Triton X-100 added to the protein before freezing (similar to the inclusion of 0.1% detergent used by Suetake et al. [43]) or to the protein 10 min before the assay, both stimulated the activity of the DNMT3A preparation 1.5- to 2-fold. If the fraction of active enzyme for DNMT3A is considered, the turnover rate constant with poly dIdC becomes 6.7 h1. This assumes that the fraction of active enzyme with poly dIdC as a substrate is the same as that determined with a 30-mer oligonucleotide (Fig. 3). This finding gives a little more credence to applying steadystate kinetics to DNMT3A here and in other studies which use poly dIdC as a substrate [43,51,55]. Multiple turnovers on poly dIdC are being observed in the typical 1 h time frame of most assays (rather than the previous one or two turnovers as previously believed). Though the total full length DNMT3A concentration in our assays (30 nM) may approach that of the lowest concentration poly dIdC substrate (300 nM in base pairs) when a 10–15 bp foot print is considered, the actual active enzyme concentration is much lower than the total DNMT3A concentration. The Km(poly dIdC) of 1.4 lM in base pairs determined here agrees with previous measurements of 1.2 lM [43] and 2.7 lM [55] for murine Dnmt3a. A higher value of 6.25 lM was reported for human DNMT3A2 [51]. The inclusion of Mg2+ in the assay buffers used for DNMT3A2, though closer to physiological conditions, could account for the difference.

DNMT3A contains a second DNA binding site Previous studies with murine Dnmt1 showed substrate inhibition by poly dIdC, plasmid, and 30mer duplex oligonucleotide substrates [10,27–29]. Further mechanistic studies utilizing a pulsechase processivity assay revealed that this inhibition was due to a second DNA molecule binding to an allosteric site of Dnmt1 [30–32]. As shown in Figs. 4a and 6, substrate inhibition was also observed for DNMT3A. In contrast to Dnmt1, the isolated catalytic domain of Dnmt3a is catalytically activity [61–63]. This fact was exploited to gain information on the mechanism of substrate inhibition of DNMT3A and to locate a region of the protein critical for binding the second molecule of DNA. The isolated catalytic domain of DNMT3A did not show substrate inhibition. This suggests that an allosteric DNA binding site found within the first 611 residues is the source of substrate inhibition, rather than the alternative mechanism of an inhibitory complex forming during product re-

lease. Two further truncations indicated that a region within residues 224–459 was necessary in forming the allosteric site. This region contains the recognized PWWP domain, which was shown to bind DNA in murine Dnmt3b [16,25] and two other proteins [24,59]. EMSA based measurements revealed that the DNMT3A PWWP domain has a DNA affinity consistent with the observed substrate inhibition phase of rate versus oligonucleotide substrate concentration plots in Figs. 5 and 6. The apparent cooperativity could result from a need for multiple PWWP domain molecules to bind to an oligonucleotide before a mobility shift is detected. Alternatively, it could indicate that the PWWP domain must aggregate in solution before it can bind to DNA. This is consistent with observed aggregation of the DNMT3A2, though the catalytic domain is believed to contain the primary DNMT3A– DNMT3A interface [51,52,60]. Aggregation or other types of instability could also simply be an artifact of working with a truncated form of the protein. Alternatively, it may suggest that the PWWP domain, while necessary for binding the second DNA molecule, might not contain the entire binding surface of the allosteric site. Other regions of the protein may be involved in either forming a portion of the allosteric binding site or in maintaining the proper conformation or stability of the PWWP domain. Similar to previous studies on the murine Dnmt3b PWWP domain [16,25] and the hepatoma-derived growth factor [24], the DNMT3A PWWP domain appears to bind DNA without sequence specificity for the few oligonucleotide duplexes tested here, including 5-methylcytosine containing duplexes and a sequence with no CpG sites. A previous single point measurement showing no DNA binding by the murine Dnmt3a PWWP domain [16] is not in direct conflict with our present measurements, given the low DNA affinity human DNMT3A PWWP domain observed here combined with the much higher ionic strength of the previous work. Potential roles for the allosteric site in DNMT3A Previous studies on the Dnmt3 PWWP domain focused on DNA binding [16,25] or on its role in targeting Dnmt3a and Dnmt3b to chromatin [16,26]. This study focuses on the kinetic consequences of DNA binding to this site. Occupancy of this second DNA binding site represses the active site turnover rate approximately 2-fold (Fig. 4a and Table 2). This could be a useful regulatory mechanism, but there is then a question of the physiological significance of a DNA binding site with weak affinity and low sequence specificity. Though most of the DNA in a mammalian nucleus would be bound to other proteins, the very high concentration (estimated to be at least 1.5 mM in base pairs [64]) suggests that the lM affinities measured here could be physiologically relevant. Four possible biological roles for this allosteric binding site are presented. The sequence space searched here was limited, suggesting that other yet to be identified sequences might impact DNMT3A’s affinity through the allosteric site or have a greater impact on its turnover rate. For Dnmt1 the potency of an inhibitor bound at its allosteric site was sequence specific and required the presence of a 5-methyl-deoxycytosine residue [29–31]. It is also possible that the DNMT3A allosteric site could specifically recognize sequences or secondary structures in single stranded DNA or RNA. Even in the absence of sequence specificity, DNA binding to DNMT3A’s PWWP domain could perform a regulatory function. A similar role has been proposed for an allosteric site of the p53 transcription factor where binding of non-specific DNA at a carboxy-terminal domain weakens sequence specific binding at the central domain of p53 [65]. In addition non-coding RNA molecules have been shown to regulate a transcription factor allosterically [34] and to facilitate targeting of the histone methylating polycomb complex [33,35]. In the case of DNMT3A, inhibition by non-specific DNA binding to the PWWP domain could enhance

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regulatory interactions with other proteins. Interactions between Dnmt3a and several other proteins have been noted [7,19–23], but the consequences of these protein–protein interactions on Dnmt3a catalysis have generally not been measured. However, thymidine DNA glycosylase binding was shown to inhibit Dnmt3a catalysis, and the Dnmt3a PWWP domain formed part of the interaction site [20]. In general, DNA non-specifically bound to DNMT3A’s PWWP domain could be displaced by protein binding in this region. Loss of inhibition could add an extra level of rate stimulation of DNMT3A by other proteins. A third possible function for a non-sequence specific allosteric DNA binding site is in modulating the enzyme’s processivity. This second binding surface could allow extra contact with a long DNA substrate, increasing the likelihood that the enzyme would stay bound to the substrate while searching for a new methylation site. As proposed for Dnmt1 [30–32], substrate inhibition could occur in this model when high concentrations of a DNA second molecule displace the portion of the substrate DNA molecule from the allosteric site. Previously, it was suggested that the full length Dnmt3a [54] and its isolated catalytic domain [61] did not methylate multi-site substrates processively. However, it is unclear in these previous studies whether or not the enzymes were undergoing the multiple turnovers required to monitor processivity. Also, the target sites chosen for analysis did not fit either of the later determined consensus sequence patterns for good Dnmt3a sites [56,66]. Effects of flanking sequence on enzymatic processivity have been observed in E. coli DNA adenine methyltransferase [44,67]. It remains possible that processivity could be observed with DNMT3A in a multiple turnover assay on poly dIdC, poly dGdC, or a substrate with multiple good sites. We have recently showed that human DNMT3A can act processively (C. Holz-Schietinger and N.O. Reich, unpublished results). Finally, it is possible that there is no regulatory or functional role for the weak DNA binding activity of this site. DNMT3A’s PWWP domain maybe primarily employed in mediating protein– protein interactions, and DNA binding may be a coincidental or vestigial activity. The PWWP domains of both Dnmt3a and Dnmt3b were shown to be critical in localizing these enzymes to chromatin, though there is disagreement as to whether a PWWP–DNA interaction makes a contribution to Dnmt3a’s chromatin association [16,26]. Conclusions An allosteric DNA binding site with weak affinity was found in DNMT3A and determined to involve the PWWP domain. Occupancy of this site lowers the rate of turnover at the active site, but the second DNA binding site appears to lack sequence specificity. Several potential roles for a low affinity, non-specific allosteric site in regulating enzyme activity were discussed. Further studies will examine the DNMT3A’s PWWP domain binding specificity in terms of sequence and type of nucleic acid (single stranded versus double stranded; DNA versus RNA) and the impact of protein–protein interactions on nucleic acid binding to this site. It will be particular interesting to see how the reorganization of DNMT3A upon pre-incubation [51] or co-expression with DNMT3L [52,60] impacts inhibition mediated through the PWWP domain. Acknowledgments The authors greatly appreciate the gifts of the cDNA clone for DNMT3A from En Li and Taiping Chen, Novartis Institutes for Biomedical Research and a synthetic codon optimized portion of the DNMT3A gene from Frédéric Chédin, University of California at

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Davis. M.M.P. was supported in part by NIH postdoctoral fellowship GM071206.

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