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Abstract

The histone methyltransferase (HMT) family of proteins consists of enzymes that methylate lysine or arginine residues on histone tails as well as other proteins. Such modifications affect chromatin structure and play a significant regulatory role in gene expression. Many HMTs have been implicated in tumorigenesis and progression of multiple malignancies and play essential roles in embryonic development and stem cell renewal. Overexpression of some HMTs has been observed and is correlated positively with various types of cancer. Here the authors report development of a continuous fluorescence-based methyltransferase assay in a 384-well format and its application in determining kinetic parameters for EHMT1, G9a, PRMT3, SETD7, and SUV39H2 as well as for screening against libraries of small molecules to identify enzyme inhibitors. They also report the development of a peptide displacement assay using fluorescence polarization in a 384-well format to assay and screen protein peptide interactions such as those of WDR5 and EED, components of MLL and EZH2 methyltransferase complexes. Using these high-throughput screening methods, the authors have identified potent inhibitors and ligands for some of these proteins.

Keywords

enzyme assays or enzyme kinetics epigenetics protein-protein interactions fluorescence methods

Introduction

Posttranslational modification of histones such as methylation, acetylation, phosphorylation, ubiquitination, sumoylation, and adenosine diphosphate (ADP)–ribosylation affect chromatin condensation and provide signals to regulate transcription, DNA replication, and repair.¹ Among these modifications, histone methylation has been widely studied. Since SUV39H1 was identified as a SET domain–containing histone methyltransferase (HMT),^2,3 many other proteins have been annotated as HMTs, which catalyze the transfer of a methyl group from S-adenosyl-L-methionine (SAM) to lysine or arginine residues on histone tails. However, many of these proteins are also involved in methylation of nonhistone proteins.^4–8 The HMT family can be clustered into smaller groups based on amino acid sequence homology⁹ (Suppl. Fig. S1).

Protein arginine methyltransferases (PRMTs) may transfer two methyl groups to a single nitrogen atom of the guanidine moiety of arginine (type I) or transfer two methyl groups to two different nitrogen atoms of the guanidine (type II).¹⁰ PRMT3 is a type I PRMT that methylates arginine 3 of histone H4.¹¹ It also methylates ribosomal protein S2 (rpS2), stabilizes it, and plays a role in proper maturation of 80S ribosome by binding to and catalyzing the methylation of rpS2.¹² Its function has been reported to be essential for dendritic spine maturation in rat.¹³ Interestingly, DAL-1/4.1B tumor suppressor interacts with PRMT3 and inhibits its HMTase activity, suggesting that the tumor suppressor may affect tumor growth through regulating protein arginine methylation.⁸ Identifying JMJD6 as a histone arginine demethylase suggests that histone arginine methylation is a dynamic process.^14,15

Lysine methyltransferases catalyze mono-, di-, and trimethylation of various histone tail lysine residues with specificity for both the degree of methylation on lysine and the surrounding amino acid sequence. SETD8 (PR-SET7) catalyzes the monomethylation of lysine 20 of histone H4 (H4K20), which is required to recruit 53BP1 to sites of DNA damage.¹⁶ ES cells lacking SETD8 display atypical chromosome condensation and massive DNA damage.¹⁷ H4K20 is not the only SETD8 substrate, as lysine 382 of tumor suppressor protein p53 is also methylated by this protein.¹⁸ SETD7 (SET7/9) is another lysine methyltransferase that methylates lysine H3K4. H3K4 methylation has been reported to impair SUV39H1-mediated methylation at lysine 9 of H3 (H3K9).¹⁹ SETD7 also monomethylates lysine 142 of DNMT1.²⁰ DNMT1 methylation has been suggested to regulate its proteasome-mediated degradation.²⁰ Methylation of TAF-10 at lysine 189²¹ and p53 at lysine 372²² by SETD7 is required for their structural and functional stabilization. Other proteins have also been reported as substrates for SETD7, including retinoblastoma tumor suppressor protein,²³ estrogen receptor α,²⁴ p65,²⁵ and Tat protein of HIV-1.⁷ SETD7 has been suggested to act as a coactivator of HIV transcription.

G9a and EHMT1 mono- and dimethylate H3K9, whereas SUV39H2 also trimethylates this residue. G9a and EHMT1 also methylate nonhistone proteins, including p53 (K372), Wiz (K305), CDYL1 (K135), ACINUS (K654),^26,27 and Reptin (K67),²⁸ as well as undergoing auto-methylation.²⁹ Methylated H3K9 is commonly associated with gene repression.^30,31 In a G9a knockout mouse, 167 genes were significantly upregulated, but no gene was downregulated, granting G9a a gatekeeper function.³² Tumor suppressor genes are among such silenced genes.³³ It has recently been reported that G9a promotes lung cancer invasion and metastasis by silencing the cell adhesion molecule Ep-CAM.³⁴ SUV39H2 polymorphisms have been shown to be associated with risk of lung cancer.³⁵

Some HMTs are only active when in complex with other proteins. Histone methyltransferase enhancer of Zeste homolog 2 (EZH2) is the catalytic subunit of polycomb repressive complex 2 (PRC2). EED and SUZ12 are essential for HMT activity of EZH2.³⁶ RbAp48 and AEBP2 may function by promoting the enzymatic activity of this complex. EZH2 catalyzes the addition of three methyl groups to histone H3K27. Trimethylation of H3K27 is responsible for the transcriptional repression of key developmental genes. EED-specific binding to trimethylated lysine residues is associated with repressive chromatin marks.³⁷ Histone methyltransferase mixed-lineage leukemia protein (MLL) is only active in complex with WDR5 and RbBP5.³⁸ WDR5, which forms a WD40 repeat β-propeller, is a key protein in the assembly of the MLL core complex.³⁹ It binds to trimethylated H3K4 as well as WDR5 interaction motif (Win) of MLL⁴⁰ and is essential for trimethylation of H3K4 by the MLL complex.^41,42

The critical roles of HMTs in a variety of diseases suggest that many of these enzymes may be targets for a new generation of therapeutics,^43,44 but the degree to which potent, selective, and nontoxic agents can be developed is still unknown. Furthermore, little is known about substrate specificity and kinetic behavior for many of the annotated HMTs. Thus, the field would benefit from the availability of a broadly applicable HMT assay to better characterize HMTs and their inhibition. In addition to inhibition of HMTase activity by compounds that bind to their active sites, HMT complexes such as PRC2 and MLL complexes can potentially be inactivated by targeting their complex formation. Compounds disrupting interaction of WDR5 with MLL and EED with EZH2 could potentially inactivate these enzymes without directly binding to their SET domains. To screen for such compounds, displacement of synthetic peptides that are similar in sequence to the interacting motifs can be used and monitored by fluorescence polarization.^45–48

Various enzyme assays have been developed during the past few years that have significantly improved our knowledge of the specificity of HMTs. A coupled fluorescent assay for histone methyltransferases was first developed as an end point assay that is applicable to most HMTs with high enough catalytic turnover.⁴⁹ In this assay, SAHH (S-adenosylhomocysteine hydrolase) and adenosine deaminase convert the methyltransferase reaction product (S-adenosyl homocysteine) to homocysteine and inosine. Homocysteine can be quantified using ThioGlo. This assay was later optimized and used as a continuous assay for screening G9a, which led to the discovery of UNC0638, a potent and selective chemical probe for G9a and EHMT1.^50–52 This continuous assay was also used to characterize nonhistone plant caffeic acid O-methyltransferases.⁵³ End point assays rely on one single readout and are prone to higher experimental error and yield higher false-positive and false-negative rates in high-throughput screening (HTS). In the continuous mode, the activity of the enzyme is calculated from the slope of signal over time plot. This way, the linearity of the initial velocity and the presence of any starting lag or leveling off within the timeframe of the assay can easily be detected. This is crucial for accurately determining the kinetic parameters for HMTs. In addition, possible effect of compounds on fluorescence readout can also be detected. Other coupled assays have also been developed using other coupling enzymes.^54,55 These methods also rely on hydrolyzing SAH to other components and monitoring such conversions. In a colorimetric assay, S-adenosylhomocysteine (SAH) is first hydrolyzed by recombinant SAH nucleosidase into adenine and S-ribosylhomocysteine. In turn, S-ribosylhomocysteine is cleaved to form homocysteine using S-ribosylhomocysteinase. Homocysteine is quantified using Ellman’s reagent, and the accompanying absorption change at 412 nm is monitored.⁵⁵ In another method, a spectrophotometric assay monitors a deamination reaction that is associated with a decrease in absorbance at 265 nm. In this assay, SAH is hydrolyzed to S-ribosylhomocysteine and adenine by recombinant S-adenosylhomocysteine/5′-methylthioadenosine nucleosidase, adenine product is further hydrolyzed to hypoxanthine and ammonia by recombinant adenine deaminase, and deamination is monitored in a continuous mode.⁵⁴

In an alternative strategy, antibodies against specific methyl-lysine histone marks have been employed in developing medium- and high-throughput assays. A dissociation-enhanced lanthanide fluoroimmunoassay (DELFIA) assay was used to screen G9a against a library of 125 000 compounds, which resulted in discovery of BIX-01294, a potent G9a inhibitor. In this assay, the biotinylated peptide containing H3K9me2 reaction product was captured by neutravidin-coated 384-well microtiter plates, and methylated peptides were detected with an H3K9me2 antibody and a europium-labeled goat α-rabbit secondary antibody.⁵⁶ Other antibody-based assays developed for HMTs include a chemiluminescence-based method⁵⁷ and an enzyme-linked immunosorbent assay (ELISA).⁵⁸ One drawback of the antibody-based detection methods is the batch-to-batch variability in quality and specificity of available antibodies. Another method uses endoproteinase-LysC to cleave the unused substrate peptide (unmethylated) followed by separation and quantification of the methylated peptide through microfluidic capillary electrophoresis using Caliper instruments. This instrument separates small biomolecules such as peptides, based on their charge-to-mass ratio. Although methylation does not alter the charge of peptide substrates, in this case, the difference in size of the cut and uncut peptides is significant enough for separation. This assay was successfully employed to assay histone methyltransferases.⁵⁹ However, it is also an end point assay. An HMTase assay monitored by the transfer of a radioactive methyl group from SAM to lysine or arginine has also been widely employed. These types of assays are very sensitive and amenable to HTS.^60,61 However, safety concerns and regulatory burdens associated with radioactivity-based assays may pose a limitation to some laboratories.

Developing high-throughput activity-based assays customized for each HMT would provide opportunities to screen each of these proteins against chemical libraries for developing potent, selective, and cell-penetrant inhibitors (chemical probes).⁵⁰ Here we report further optimization and validation of an SAHH-coupled assay in a continuous mode to determine kinetic parameters and identify screening conditions for G9a, EHMT1, SUV39H2, SETD7, and PRMT3, as well as an end point assay for SETD8 in a 384-well format. We also report the optimization and validation of a peptide displacement assay for screening WDR5 and EED.

Material and Methods

Adenosine deaminase (ADA) was purchased from Sigma (cat. 096K7003; Sigma, St. Louis, MO), ThioGlo from Calbiochem (cat. 595501; Calbiochem, Gibbstown, NJ), and SAH from Sigma (cat. A9384). 384-well plates were purchased from Axygen (cat. PCR-384-BK; Axygen, Union City, CA), and Greiner (cat. 784209; Greiner Bio-One, Monroe, NC). The 96-well plates were obtained from Nalgene (cat. 249944; Nalgene, Rochester, NY).

Human HMTs used in this study were cloned, expressed, and purified as described before.⁶² The SsSAHH clone was provided by Dr. Raymond Trievel (University of Michigan). The ApSAHH clone was provided by Dr. Alexei Savchenko (University of Toronto).

SAHH Assay

In this assay, SAHH and ADA convert SAH to homocysteine and inosine. The abundance of homocysteine was quantified using ThioGlo, which reacts with thiol groups and fluoresces strongly. To obtain the K_m and k_cat values for SAH, a SAH concentration ranging from 10 nM to 50 µM in a series of experiments was mixed with ADA (final 0.5 U/mg) in 25 mM potassium phosphate (pH 8), 1 mM EDTA, 2 mM MgCl₂, 0.01% Triton X-100, ThioGlo at a final concentration of 15 µM, and SAHH at a final concentration of 1 µM in a 20-µL reaction mix. Then, 9 µL of reaction mix, in duplicate, was transferred into each well of a 384-well plate (Axygen). The conversion of SAH to homocysteine was followed by monitoring the increase in fluorescence using a ViewLux (PerkinElmer, Waltham, MA) plate reader with 340 DUG 11 filter for excitation and 540/25 filter for emission for 20 min in a 384-well plate format. Homocysteine generated in the assay was quantified using standard curves. Activity values were corrected by subtracting background caused by SAH. K_m values were calculated using a hyperbolic function (Sigma Plot software, Systat Software, Chicago, IL). Standard deviations were calculated from two or more independent experiments.

HMT Assay

The methyltransferase activities of HMTs were measured using a coupled assay.⁴⁹ In this assay, SAHH and ADA convert the methyltransferase reaction product SAH to homocysteine and inosine. The abundance of homocysteine was quantified using ThioGlo. For the K_m and k_cat determination, each protein concentration was selected to be within the linear range. For G9a, EHMT1, SETD7, SETD8, SUV39H2, and PRMT3, the concentrations used were 50 nM, 100 nM, 500 nM, 200 nM, 100 nM, and 500 nM, respectively. The rate of product formation for these HMTs was constant, and the plot of signal versus time was linear over the assay period (Suppl. Fig. S2). No lag at the start of the reactions was observed. The peptides used in this assay varied for each enzyme. Thus, for G9a, EHMT1, SUV39H2, and SETD7, the substrate is the first 25 residues of histone 3 (H3 [1–25], ARTKQTARKSTGGKAPRKQ LATKAA). The substrate for PRMT3 and SETD8 is the first 24 residues of histone 4 (H4 [1–24], SGRGKGGKGLG KGGAKRHRKVLRD). For the determination of K_m values of SAM and peptide, peptide or SAM was titrated into the assay mixture containing enzyme, 5 µM SAHH, about 0.5 U/mL ADA, and 15 µM ThioGlo in 25 mM potassium phosphate (pH 8), 1 mM EDTA, 2 mM MgCl₂, and 0.01% Triton X-100. A saturating concentration of the other substrate (fixed concentration of SAM or peptide) was added to start a reaction. K_m values were calculated using Sigma Plot (Systat Software). The methylation reaction was followed by monitoring the increase in fluorescence using a ViewLux (PerkinElmer) plate reader with 340 DUG 11 filter for excitation and 540/25 filter for emission or Synergy 2 microplate reader (BioTek, Winooski, VT) with a 360/40-nm excitation filter and 528/20-nm emission filter for 20 min in a 384-well plate format. Homocysteine generated in the assay was quantified using standard curves. Activity values were corrected by subtracting background caused by the peptide and the protein. ThioGlo has an inhibitory effect on SETD8 activity due to a reactive cysteine residue near the active site. Therefore, this enzyme was assayed using the end point assay. Reactions mixtures (all components of the assay) were incubated for 4 min before adding ThioGlo at a final concentration of 15 µM and quantifying the amount of produced SAH. For assays performed over a longer period of time, a decrease in protein concentration may be required to maintain linearity of the reaction.

Z Factor Determination

Z factors were determined to evaluate the suitability of each screen in a 384-well format.⁶³ To calculate the Z factor of the assay, the HMT reactions (for each enzyme separately; see above for details) were run in eight complete rows (half-plate) of 16 × 24 of a 384-well plate; the other eight rows were used for background controls (the reaction mix containing protein but not SAM). The reaction volume for G9a, EHMT1, SUV39H2, and PRMT3 was 10 µL. For SETD7 and SETD8, the reaction volume was increased to 20 µL to obtain a better signal-to-noise ratio. To obtain a Z′ factor value for the assay, a known inhibitor (UNC-0638 in case of G9a) was added at 1 µM to the reaction mix. The Z factor for each protein was calculated as described by Zhang et al.⁶³

Fluorescence Polarization

H3K27me3 (QLATKAARK(me3)SAPATG) peptide for EED and MLL_Win (GSARAEVHLRKS) peptide for WDR5 were synthesized, N-terminal-labeled with isothiocyanate–fluorescein, and purified by Tufts University Core Services (Boston, MA). Compound binding assays were performed in a 10-µL volume at a constant labeled peptide concentration of 40 nM and protein concentrations of 80 µM for EED and 5 µM for WDR5. For EED, the buffer contained 20 mM Tris (pH 8.0), 50 mM NaCl, 2 mM dithiothreitol (DTT), and 0.01% Triton X-100. For WDR5, the buffer was 100 mM potassium phosphate (pH 8.0), 150 mM NaCl, and 0.01% Triton X-100. Fluorescence polarization assays were performed in 384-well Axygen plates using a Synergy 2 microplate reader (BioTek). The excitation wavelength of 485 nm and the emission wavelength of 528 nm were used. To determine K_d values, the data were fit to a hyperbolic function using Sigma Plot software (Systat Software). The K_d values represent the average and standard deviation of three measurements.

Results and Discussion

Assay and Screening Writers of Histone Methyl Markers

The SAHH-coupled assay developed by Collazo et al.⁴⁹ is a convenient method to assay a wide range of HMTs. In this assay, the HMTase activity is coupled with SAHH activity. This coupled assay relies on SAHH using SAH, the product of the HMTase reaction as a substrate and producing L-homocysteine and adenosine. The free SH group of homocysteine then reacts with ThioGlo, and the fluorescence signal will increase ( Fig. 1 ). Such an increase in fluorescence signal reflects HMTase activity, and the amount of produced SAH can be quantified. In this study, we have optimized this assay in a 384-well format as a continuous assay and determined the kinetic parameters for G9a, EHMT1, SUV39H2, SETD7, and PRMT3. SETD8, however, was not amenable to the continuous assay and was characterized using the end point assay.

Figure 1.

S-adenosylhomocysteine hydrolase (SAHH) assay using fluorescence detection. L-homocysteine, the product of SAHH function, reacts with ThioGlo, and emission at 513 nm (excitation of 384 nm) can be monitored to detect S-adenosylhomocysteine (SAH) production and therefore SAHH activity. Presence of adenosine deaminase takes away adenosine, the by-product of the SAHH reaction, by converting it to inosine to prevent possible reverse reaction

Assay Optimization

Characterization of SAHH at 28 °C from Sulfolobus solfataricus (SsSAHH)⁴⁹ with a K_m of 1.8 ± 0.1 (µM) for SAH and k_cat of 20 ± 0.3 h⁻¹ and Aeropyrum pernix (ApSAHH) with a K_m of 1.2 ± 0.1 (µM) and k_cat of 21 ± 0.1 h⁻¹ ( Table 1 ) indicated that these two proteins have more or less similar kinetic parameters, with ApSAHH having slightly higher specific activity (18 h⁻¹ µM⁻¹) compared to that for SsSAHH (11 h⁻¹ µM⁻¹). However, we selected SsSAHH for assay optimization because all preliminary experiments were previously performed using this SAHH.

Table 1.

Characterization of SsSAHH and ApSAHH

Protein	Source	K_m(µM)	k_cat(h⁻¹)	k_cat/K_m (h⁻¹ µM⁻¹)
SsSAHH	Sulfolobus solfataricus	1.8 ± 0.1	20 ± 0.3	11
ApSAHH	Aeropyrum pernix	1.2 ± 0.1	21 ± 0.1	18

Collazo et al.⁴⁹ used an end point assay to determine kinetic parameters of HMTs by initiating the reaction with addition of a saturating concentration of SAM in a large volume and aliquoting the reaction mix and stopping the reaction at evenly spaced time intervals starting at time zero by cold isopropanol. After the reaction was completed, a solution of ThioGlo in DMSO was added to each well and allowed to react for 10 min at room temperature, and fluorescence was then measured. To determine the quantity of homocysteine generated in the assay, reduced CoA was used to generate standard curves.

Here we modified the reaction so that we can monitor the production of SAH (through detecting homocysteine) continuously. A reaction mix was prepared with all components of the assay, including protein and SAM but not peptide. Freshly prepared ThioGlo was added to this reaction mix along with SAHH and ADA. The reaction was started by adding the respective peptide substrate for each protein, and the increase in SAH production was monitored by recording the emission at 528 nm (excitation at 360 nm) using Synergy2, Synergy4, or ViewLux plate readers. It was crucial to prepare the ThioGlo solution freshly each time to obtain reproducible data. The ThioGlo should be stored in 100% DMSO and then added to buffer assays, which are slightly alkaline (pH 8). To ensure that the SAHH reaction is not a rate-limiting step and SAH conversion to homocysteine is properly monitored, we performed the assay for G9a, the most active HMT in our set, at saturating concentrations of SAM and H3 (1–25) peptide and varying the SsSAHH concentration ( Fig. 2A ). The fluorescence signal increased by increasing the SAHH up to 4 µM and plateaued completely at a 5-µM concentration. To ensure that conversion by SAHH is not a rate-limiting step, we also tested the activity of the HMTs using 5 and 10 µM SAHH. Doubling the concentration of SAHH made no significant difference (Suppl. Fig. S3). Therefore, we selected the 5-µM concentrations of SAHH for all following experiments in our HMT set. Another component of the coupled assay to consider was ADA. This enzyme was originally included in this assay to take away adenosine, the by-product of the SAHH reaction ( Fig. 1 ), to reduce the possibility of reverse reaction and SAH production upon SAHH-catalyzed condensation of adenosine and homocysteine.⁴⁹ However, it was later reported that ADA may not be required in the plant caffeic acid O-methyltransferases assay.⁵³ We also performed the SAHH-coupled assay for G9a at 5 µM SsSAHH and varying the concentration of ADA ( Fig. 2B ). No significant effect of ADA concentration on activity of SAHH was observed. However, due to a small increase in the level of activity at low concentrations of ADA ( Fig. 2B ), we selected to use 5 µM SAHH and 0.5 U/mL ADA in all HMT assays. The concentration of HMTs in the assay must be kept low enough so that all produced SAH is simultaneously converted to homocysteine and subsequently reacts with ThioGlo. The SAHH reaction should not be a rate-limiting step at any point in the continuous assay. Therefore, we performed all the assays at 5 µM SAHH and around 50, 100, 100, 500, and 500 nM of G9a, EHMT1, SUV39H2, SETD7, and PRMT3, respectively.

Figure 2.

Effect of S-adenosylhomocysteine hydrolase (SAHH) and adenosine deaminase (ADA) on histone methyltransferase (HMT) assay readout. G9a activity was assayed at 0.5 U/mL ADA and different concentrations of SAHH (A) and at 5 µM SAHH and different concentrations of ADA (B). The buffer was 25 mM potassium phosphate (pH 8), 1 mM EDTA, 2 mM MgCl₂, and 0.01% Triton X-100. Reaction mix contained saturating concentrations of S-adenosyl-L-methionine (SAM; 10 µM) and peptide (10 µM) substrate (H3 [1–25]). Experiments were performed at 28 °C

Determining the Kinetic Parameters for HMTs

We employed this continuous assay to characterize G9a, EMHT1, SUV39H2, SETD7, PRMT3, and SETD8. However, due to interaction of ThioGlo with a free cysteine thiol in SETD8 and a clearly related inhibitory effect, we excluded SETD8 from this set and determined its kinetic parameters using the end point assay.⁴⁹ Kinetic parameters for all these HMTs are summarized in Table 2 . G9a, EHMT1, and SUV39H2 methylate lysine 9 (H3K9), and SETD7 methylates lysine 4 (H3K4) of histone H3. We performed all kinetic assays for these four HMTs using a 25-mer peptide corresponding to the first 25 residues of histone H3 (H3 1–25) as the substrate. SETD8, on the other hand, methylates lysine 20 of histone H4, and PRMT3, being an arginine methyltransferase, methylates arginine 3 of histone H4 (H4R3). A peptide corresponding to the first 24 residues of histone H4 was used as a substrate for these two proteins. To facilitate purification of all these HMTs, we used shorter active constructs of these proteins ( Table 2 ). For the three H3K9 HMTs, the K_m values for the peptide substrate (H3 1–25) are more or less in the same range (1.7–3.4 µM). However, the size of the peptide substrate significantly can affect the K_m of the protein for the peptide. For example, with G9a, the K_m for H3 (1–11) was determined to be 50 ± 7 µM, almost 30 times higher than that for H3 (1–25). Among these HMTs, G9a showed the highest specific activity (450 h⁻¹ µM⁻¹). Reproducibility of determining the kinetic parameters for these proteins confirmed the suitability of this assay for characterizing most HMTs.

Table 2.

Kinetic Parameters of HMTs

				K_m (µM)
Protein	Construct	Number of Amino Acids	Peptide Substrate	Peptide	SAM	k_cat (h⁻¹)	k_cat/K_m (h⁻¹ µM⁻¹)
G9a	913–1193	281	H3 (1–25)	1.7 ± 0.1	1.5 ± 0.1	765 ± 8	450
EHMT1	951–1235	253	H3 (1–25)	3.4 ± 0.1	1.2 ± 0.2	465 ± 30	136
SUV39H2	52–350	299	H3 (1–25)	3 ± 0.1	8 ± 0.1	660 ± 5	220
SETD7	1–366	366	H3 (1–25)	160 ± 40	0.6 ± 0.2	25 ± 2	0.15
SETD8	186–352	167	H4 (1–24)	74 ± 14	38 ± 6	960 ± 40	12
PRMT3	211–531	230	H4 (1–24)	13 ± 3	25 ± 5	50 ± 4	4

HMT, histone methyltransferase; SAM, S-adenosyl-L-methionine.

Among H3K9 methyltransferases studied here, G9a and EHMT1 mono- and dimethylate lysine 9. SUV39H2, however, also trimethylates H3K9.⁶² Our data indicate that the K_m of the peptide increases for higher methylation states (un- < mono- < dimethylated H3 [1–25]; Table 3 ). With k_cat values slightly increasing for higher methylation states, SUV39H2 shows much higher specific activity with unmethylated peptide.

Table 3.

Substrate Specificity of SUV39H2

		K_m (µM)
Protein	Peptide Substrate	Peptide	SAM	k_cat (h⁻¹)	k_cat/K_m (h⁻¹ µM⁻¹)
SUV39H2	H3 (1–25)	3 ± 0.1	8 ± 0.1	660 ± 5	220
SUV39H2	H3 (1–25) K9Me1	12 ± 2	ND	860 ± 55	71
SUV39H2	H3 (1–25) K9Me2	60 ± 8	ND	940 ± 110	15

SAM, S-adenosyl-L-methionine; ND, not determined.

HTS Using Continuous Assay

HMTs methylate histones and affect chromatin condensation and gene expression. HMTs have been implicated in many diseases, including various types of cancer.^1,34,64 Identifying potent inhibitors of HMTs with in vivo activity could be very useful to further investigate such effects and may provide the basis for drug discovery. Two sets of potential inhibitors can be identified for HMTs: (1) peptide mimetic compounds that bind to the peptide binding site and therefore compete with peptide such as the inhibitors reported for G9a^50–52,56 and (2) compounds that may bind to SAM binding site and compete with SAM. To identify peptide competitive inhibitors, one needs to screen HMTs against libraries of compounds at low concentrations of peptide substrates while keeping the concentration of SAM close to saturation. Typically, screening at a peptide concentration close to the K_m of peptide and a SAM concentration about five times the K_m of SAM should provide an opportunity for compounds to compete with the peptide for binding to the peptide binding site and make hit detection easier. We assessed such screening in a 384-well format for all five HMTs in this study, and Z factors of 0.64, 0.76, 0.77, 0.84, and 0.84 were obtained for SETD7, G9a, EHMT1, SUV39H2, and PRMT3, respectively, indicating that this continuous assay is very suitable for HTS (Suppl. Table S1 and Fig. 3A ). We employed this method to screen SUV39H2 and PRMT3 against a library of thousands of diverse compounds, and we identified hits with low micromolar IC₅₀ values (e.g., PDB ID: 3SMQ). A Z factor of 0.84 was also determined for screening SETD8 at a K_m of peptide using the end point SAHH-coupled assay.

Figure 3.

Screening G9a in a 384-well format. G9a was screened at peptide concentration close to the K_m for peptide (2 µM) and saturation concentration (10 µM) of S-adenosyl-L-methionine (SAM; A) and SAM concentration close to K_m of SAM (2 µM) and saturation concentration (10 µM) of H3 (1–25) peptide (B) to determine Z factors. G9a was also screened in the presence (185 samples) and absence (185 samples) of UNC0638 (a potent G9a inhibitor) at conditions similar to (A), and a Z′ factor of 0.71 was obtained

On the other hand, all these HMTs use SAM as a cofactor, and identifying compounds that can compete with SAM, yet are selective enough among HMTs, may seem more difficult. Through a systematic analysis of SAM binding site across structures of 14 human HMTs or close homologues, it was shown that SAM binding pockets are likely diverse enough to be chemically tractable, and identifying selective SAM competitive inhibitors is possible.⁹ To identify SAM competitive inhibitors, we also assessed screening HMTs at the K_m of SAM and about five times the K_m of peptides. Generally, at this condition, less variation was observed and better Z factors were obtained (Suppl. Table S1 and Fig. 3B ). Alternatively, if one is not looking for a specific type of inhibitor, screening HMTs at SAM and peptide concentrations close to respected K_m values will allow detection of any type of inhibitors. Further kinetic analysis could determine the mechanism of inhibition. However, by lowering both SAM and peptide concentrations, the signal and reproducibility may decrease, resulting in a high number of false positives.

To see how reproducibly a potent inhibitor can be identified through such screening, G9a was screened in the presence and absence of UNC0638.⁵⁰ Screening was reproducible in a 384-well format with a Z′ factor of 0.71 ( Fig. 3C ). Our results show that this assay can reliably be used to screen HMTs against libraries of a large number of compounds. However, in such screening, compounds that inhibit SAHH activity or quenching the fluorescence signal may also be identified as HMT inhibitors (screening hits). Therefore, the screening process should include a counterscreen to eliminate false positives. The SAHH activity assay (using SAH as a substrate for SAHH) is a suitable counterscreen that includes all assay components. HMT screening hits that also reduce the signal in this counterscreen are false positives. Although a continuous SAHH-coupled assay is perfectly suitable for characterization of HMTs and HTS, confirmation of a high number of screening hits by other high-throughput orthogonal methods, such as recently developed chemiluminescence,⁵⁷ microfluidic capillary electrophoresis,⁵⁹ or radioactivity-based assays, may be more convenient before proceeding with further characterization using low-throughput methods (Isothermal titration calorimetry; ITC, etc.) or follow-up chemistry. The SAHH-coupled assay is an easy to optimize assay for screening other HMTs.

Screening Histone Methyl Marks Using Peptide Displacement

Enzymatic activity of some methyltransferases such as MLL and EZH2 depends on their interaction with other proteins. MLL catalyzes histone H3 lysine 4 methylation, and its enzymatic activity depends on interaction with a three-subunit complex composed of WDR5, RbBP5, and Ash2L.⁶⁵ PRC2 complex includes EED and SUZ12 components, which interact with EZH2. This complex methylates histone H3K27 and mediates gene silencing. PRC2 components are overexpressed in metastatic prostate cancer.⁶⁶ As these components seem to be essential for methyltransferase activity of these complexes, we developed an assay to screen the reader components of these methyltransferase complexes.

WDR5 binds to MLL through recognition of a specific motif (WIN motif) of MLL, which includes an arginine residue (3765), which binds to the same arginine binding pocket on WDR5 that has been shown to bind histone H3.⁴⁰ Any compound that interrupts interaction of MLL and WDR5 may potentially inhibit MLL activity. We used a fluorescein-labeled peptide (F-peptide) (GSARAEVHLRKS) corresponding to MLL WIN motif to create a fluorescence-polarized signal ( Fig. 4A ). F-peptide (40 nM) was almost saturated at 5 µM of WDR5, with a K_d value of 1.85 ± 0.35 µM, and created a strong enough signal to assess binding of possible ligands. To test if displacement of the F-peptide can be detected and monitored upon ligand binding, we used unlabeled WIN peptide to displace the labeled WIN peptide ( Fig. 4B ). F-labeled peptide was displaced by unlabeled peptide with a K_displacement (K_dis) value of 7 ± 3.4 µM. Because interaction of WDR5 with WIN peptide was strong enough to generate a reliable signal-to-noise ratio at low WDR5 concentration (5 µM), we proceeded with testing the possibility of screening WDR5 in a 384-well format. Fluorescence polarization (FP) signal was measured in the presence of F-peptide only, in 176 wells, and by addition of 25 µM of unlabeled peptide to 176 other sample wells at 5 µM of WDR5 ( Fig. 4C ). Displacement of F-peptide by unlabeled WIN peptide was clear and detectable with a Z′ factor of 0.6. We applied this method to screen WDR5 against the above-mentioned diverse library of compounds and identified hits with K_dis of 1 to 10 µM. Binding of these hits was also confirmed by ITC (e.g., PDB ID: 3SMR).

Figure 4.

Screening WDR5 for ligand binding by monitoring peptide displacement. Binding of a fluorescein-labeled WIN peptide (GSARAEVHLRKS; 40 nM) to an increasing amount of WDR5 generated a fluorescence-polarized (FP) signal (A). Fluorescein-labeled peptide was displaced by increasing the amount of unlabeled WIN peptide (B), resulting in a decrease in FP signal. FP signal was also measured in the presence (25 µM; 176 samples) and absence (176 samples) of unlabeled peptide in a 384-well format using 5 µM WDR5 (C), and a Z′ factor of 0.6 was determined. Experiments were performed in triplicate

EED is the reader component of the EZH2 complex and binds to H3K27me3 and may be responsible for spreading this mark along chromatin.⁶⁷ Here we used a similar approach to assess ligand binding to EED. Binding of a fluorescein-labeled H3K27me3 peptide (QLATKAARKme3SAPATG; 40 nM) to an increasing amount of EED generated an FP signal ( Fig. 5A ). Fluorescein-labeled peptide was then displaced by an increasing amount of unlabeled H3K27me3 peptide ( Fig. 5B ), resulting in an FP signal decrease. Unlabeled H3K27me3 peptide displaced the labeled peptide with K_dis of 150 ± 25 µM. Although saturation was not reached up to 100 µM EED, using a 384-well format, we were able to screen EED using peptide displacement with a Z′ factor of 0.5 ( Fig. 5C ).

Figure 5.

Screening EED for ligand binding by monitoring peptide displacement. Binding of a fluorescein-labeled H3K27me3 peptide (QLATKAARKme3SAPATG; 40 nM) to an increasing amount of EED generated a fluorescence-polarized (FP) signal (A). Fluorescein-labeled peptide was displaced by an increasing amount of unlabeled H3K27me3 peptide (B), resulting in a decrease in FP signal. FP signal was measured in the presence (1 mM; 185 samples) and absence (185 samples) of unlabeled peptide in a 384-well format using 80 µM EED (C), and a Z′ factor of 0.5 was determined. Experiments were performed in triplicate

The kinetic parameters previously reported for the proteins characterized in this study are listed in the Supplementary Table S2.^43,68–74 These kinetic parameters were determined by variety of methods (radioactivity, fluorescence, mass spectrometry–based enzyme assays, etc.), each with certain assay limitations. Although the kinetic values reported here ( Table 2 ) are reasonably close to those reported before (Suppl. Table S2), in most cases, using different protein constructs (size and source, etc.), substrates (peptides with different length and sequence or histones as substrate), tags used to express and purify these proteins, and different assay conditions (pH, salt, temperature, etc.) often makes direct comparison of kinetic values quite difficult. For example, we observed significant differences in kinetic parameters for G9a when a different length of histone peptide was used.

In conclusion, the SAHH-coupled assay in a continuous mode is a reliable method for characterizing and screening G9a, EHMT1, SUV39H2, SETD7, and PRMT3 and probably many other HMTs. However, the amenability of other HMTs to this assay needs to be assessed before using this method for their characterization or screening as proteins such as SETD8 may react with ThioGlo and lose activity. In this report, we provided the kinetic parameters and optimized conditions for screening these HMTs. These kinetic values may vary for peptides with different amino acid sequences, and using nucleosomes as a substrate may produce significantly different values depending on the mechanism of substrate recognition. However, using peptides as substrates for HTS is often more convenient. Optimized screening conditions for WDR5 and EED also present a reliable method to set up screening conditions for other readers of histone methyl marks and peptide binding proteins. Overall, these assays are reliable screening methods and relatively easy to optimize for other writers and readers of histone methyl marks.

Footnotes

Acknowledgements

We thank Dr. Raymond Trievel (University of Michigan) for providing SsSAHH clone and Drs. Alexei Savchenko and Craig Daniels (University of Toronto) for ApSAHH. The Structural Genomics Consortium is a registered charity (number 1097737) that receives funds from the Canadian Institutes for Health Research, the Canadian Foundation for Innovation, Genome Canada through the Ontario Genomics Institute, GlaxoSmithKline, Karolinska Institutet, the Knut and Alice Wallenberg Foundation, the Ontario Innovation Trust, the Ontario Ministry for Research and Innovation, Merck & Co., the Novartis Research Foundation, the Swedish Agency for Innovation Systems, the Swedish Foundation for Strategic Research, and the Wellcome Trust.

Supplementary material for this article is available on the Journal of Biomolecular Screening Web site at .

References

Kouzarides

Chromatin Modifications and Their Function. Cell 2007, 128(4), 693–705.

O’Carroll

Scherthan

Peters

A. H.

Opravil

Haynes

A. R.

Laible

Rea

Schmid

Lebersorger

Jerratsch

. Isolation and Characterization of Suv39h2, a Second Histone H3 Methyltransferase Gene That Displays Testis-Specific Expression. Mol. Cell. Biol. 2000, 20(24), 9423–9433.

Rea

Eisenhaber

O’Carroll

Strahl

B. D.

Sun

Z. W.

Schmid

Opravil

Mechtler

Ponting

C. P.

Allis

C. D.

. Regulation of Chromatin Structure by Site-Specific Histone H3 Methyltransferases. Nature 2000, 406(6796), 593–599.

Dhayalan

Kudithipudi

Rathert

Jeltsch

Specificity Analysis-Based Identification of New Methylation Targets of the SET7/9 Protein Lysine Methyltransferase. Chem. Biol. 2011, 18(1), 111–120.

Huang

Dorsey

Chuikov

Perez-Burgos

Zhang

Jenuwein

Reinberg

Berger

S. L.

G9a and Glp Methylate Lysine 373 in the Tumor Suppressor p53. J. Biol. Chem. 2010, 285(13), 9636–9641.

Liu

Wang

Zhao

Zheng

Wang

Roeder

R. G.

Zhu

W. G.

Methyltransferase SET7/9 Regulates p53 Activity by Interacting with Sirtuin 1 (Sirt1). Proc. Natl. Acad. Sci. U. S. A. 2011, 108(5), 1925–1930.

Pagans

Kauder

S. E.

Kaehlcke

Sakane

Schroeder

Dormeyer

Trievel

R. C.

Verdin

Schnolzer

Ott

The Cellular Lysine Methyltransferase SET7/9-KMT7 Binds HIV-1 TAR RNA, Monomethylates the Viral Transactivator Tat, and Enhances HIV Transcription. Cell Host Microbe 2010, 7(3), 234–244.

Singh

Miranda

T. B.

Jiang

Frankel

Roemer

M. E.

Robb

V. A.

Gutmann

D. H.

Herschman

H. R.

Clarke

Newsham

I. F.

Dal-1/4.1b Tumor Suppressor Interacts with Protein Arginine N-Methyltransferase 3 (PRMT3) and Inhibits Its Ability to Methylate Substrates In Vitro and In Vivo. Oncogene 2004, 23(47), 7761–7771.

Campagna-Slater

Mok

M. W.

Nguyen

K. T.

Feher

Najmanovich

Schapira

Structural Chemistry of the Histone Methyltransferases Cofactor Binding Site. J. Chem. Inf. Model 2011, 51(3), 612–623.

10.

Bedford

M. T.

Richard

Arginine Methylation an Emerging Regulator of Protein Function. Mol. Cell 2005, 18(3), 263–72.

11.

Tang

Gary

J. D.

Clarke

Herschman

H. R.

PRMT 3, a Type I Protein Arginine N-Methyltransferase That Differs from PRMT1 in Its Oligomerization, Subcellular Localization, Substrate Specificity, and Regulation. J. Biol. Chem. 1998, 273(27), 16935–16945.

12.

Bachand

Silver

P. A.

PRMT3 Is a Ribosomal Protein Methyltransferase That Affects the Cellular Levels of Ribosomal Subunits. Embo. J. 2004, 23(13), 2641–2650.

13.

Miyata

Mori

Tohyama

PRMT3 Is Essential for Dendritic Spine Maturation in Rat Hippocampal Neurons. Brain Res. 2010, 1352, 11–20.

14.

Chang

Chen

Zhao

Bruick

R. K.

JMJD6 Is a Histone Arginine Demethylase. Science 2007, 318(5849), 444–447.

15.

Pal

Sif

Interplay between Chromatin Remodelers and Protein Arginine Methyltransferases. J. Cell Physiol. 2007, 213(2), 306–315.

16.

Oda

Hubner

M. R.

Beck

D. B.

Vermeulen

Hurwitz

Spector

D. L.

Reinberg

Regulation of the Histone H4 Monomethylase PR-Set7 by CRL4(Cdt2)-Mediated PCNA-Dependent Degradation during DNA Damage. Mol. Cell 2010, 40(3), 364–376.

17.

Oda

Okamoto

Murphy

Chu

Price

S. M.

Shen

M. M.

Torres-Padilla

M. E.

Heard

Reinberg

Monomethylation of Histone H4-Lysine 20 Is Involved in Chromosome Structure and Stability and Is Essential for Mouse Development. Mol. Cell Biol. 2009, 29(8), 2278–2295.

18.

Shi

Kachirskaia

Yamaguchi

West

L. E.

Wen

Wang

E. W.

Dutta

Appella

Gozani

Modulation of p53 Function by Set8-Mediated Methylation at Lysine 382. Mol. Cell 2007, 27(4), 636–646.

19.

Nishioka

Chuikov

Sarma

Erdjument-Bromage

Allis

C. D.

Tempst

Reinberg

Set9, a Novel Histone H3 Methyltransferase That Facilitates Transcription by Precluding Histone Tail Modifications Required for Heterochromatin Formation. Genes Dev. 2002, 16(4), 479–489.

20.

Esteve

P. O.

Chin

H. G.

Benner

Feehery

G. R.

Samaranayake

Horwitz

G. A.

Jacobsen

S. E.

Pradhan

Regulation of DNMT1 Stability through SET7-Mediated Lysine Methylation in Mammalian Cells. Proc. Natl. Acad. Sci. U. S. A. 2009, 106(13), 5076–5081.

21.

Del Rizzo

P. A.

Couture

J. F.

Dirk

L. M.

Strunk

B. S.

Roiko

M. S.

Brunzelle

J. S.

Houtz

R. L.

Trievel

R. C.

Set7/9 Catalytic Mutants Reveal the Role of Active Site Water Molecules in Lysine Multiple Methylation. J. Biol. Chem. 2010, 285(41), 31849–31858.

22.

Kurash

J. K.

Lei

Shen

Marston

W. L.

Granda

B. W.

Fan

Wall

Gaudet

Methylation of p53 by SET7/9 Mediates p53 Acetylation and Activity In Vivo. Mol. Cell 2008, 29(3), 392–400.

23.

Munro

Khaire

Inche

Carr

La Thangue

N. B.

Lysine Methylation Regulates the PRB Tumour Suppressor Protein. Oncogene 2010, 29(16), 2357–2367.

24.

Subramanian

Jia

Kapoor-Vazirani

Powell

D. R.

Collins

R. E.

Sharma

Peng

Cheng

Vertino

P. M.

Regulation of Estrogen Receptor Alpha by the SET7 Lysine Methyltransferase. Mol. Cell 2008, 30(3), 336–347.

25.

C. K.

Baltimore

Regulation of NF-kappaB Activity through Lysine Monomethylation of p65. Proc. Natl. Acad. Sci. U. S. A. 2009, 106(45), 18972–18977.

26.

Rathert

Dhayalan

Murakami

Zhang

Tamas

Jurkowska

Komatsu

Shinkai

Cheng

Jeltsch

Protein Lysine Methyltransferase G9a Acts on Non-Histone Targets. Nat. Chem. Biol. 2008, 4(6), 344–346.

27.

Huang

Berger

S. L.

The Emerging Field of Dynamic Lysine Methylation of Non-Histone Proteins. Curr. Opin. Genet. Dev. 2008, 18(2), 152–158.

28.

Lee

J. S.

Kim

I. S.

Kim

Choi

H. J.

Lee

J. M.

Shin

H. J.

Kim

J. H.

Kim

J. Y.

Seo

S. B.

. Negative Regulation of Hypoxic Responses via Induced Reptin Methylation. Mol. Cell 2010, 39(1), 71–85.

29.

Sampath

S. C.

Marazzi

Yap

K. L.

Sampath

S. C.

Krutchinsky

A. N.

Mecklenbrauker

Viale

Rudensky

Zhou

M. M.

Chait

B. T.

. Methylation of a Histone Mimic within the Histone Methyltransferase G9a Regulates Protein Complex Assembly. Mol. Cell 2007, 27(4), 596–608.

30.

Tachibana

Ueda

Fukuda

Takeda

Ohta

Iwanari

Sakihama

Kodama

Hamakubo

Shinkai

Histone Methyltransferases G9a and GLP Form Heteromeric Complexes and Are Both Crucial for Methylation of Euchromatin at H3-K9. Genes Dev. 2005, 19(7), 815–826.

31.

Tachibana

Sugimoto

Nozaki

Ueda

Ohta

Ohki

Fukuda

Takeda

Niida

Kato

. G9a Histone Methyltransferase Plays a Dominant Role in Euchromatic Histone H3 Lysine 9 Methylation and Is Essential for Early Embryogenesis. Genes Dev. 2002, 16(14), 1779–1791.

32.

Yokochi

Poduch

Ryba

Hiratani

Tachibana

Shinkai

Gilbert

D. M.

G9a Selectively Represses a Class of Late-Replicating Genes at the Nuclear Periphery. Proc. Natl. Acad. Sci. U. S. A. 2009, 106(46), 19363–19368.

33.

Wozniak

R. J.

Klimecki

W. T.

Lau

S. S.

Feinstein

Futscher

B. W.

5-Aza-2′-Deoxycytidine-Mediated Reductions in G9a Histone Methyltransferase and Histone H3 K9 Di-Methylation Levels Are Linked to Tumor Suppressor Gene Reactivation. Oncogene 2007, 26(1), 77–90.

34.

Chen

M. W.

Hua

K. T.

Kao

H. J.

Chi

C. C.

Wei

L. H.

Johansson

Shiah

S. G.

Chen

P. S.

Jeng

Y. M.

Cheng

T. Y.

. H3K9 Histone Methyltransferase G9a Promotes Lung Cancer Invasion and Metastasis by Silencing the Cell Adhesion Molecule Ep-CAM. Cancer Res. 2010, 70(20), 7830–7840.

35.

Yoon

K. A.

Hwangbo

Kim

I. J.

Park

Kim

H. S.

Kee

H. J.

Lee

J. E.

Jang

Y. K.

Park

J. G.

Lee

J. S.

Novel Polymorphisms in the SUV39H2 Histone Methyltransferase and the Risk of Lung Cancer. Carcinogenesis 2006, 27(11), 2217–2222.

36.

Kuzmichev

Margueron

Vaquero

Preissner

T. S.

Scher

Kirmizis

Ouyang

Brockdorff

Abate-Shen

Farnham

. Composition and Histone Substrates of Polycomb Repressive Group Complexes Change during cellular differentiation. Proc. Natl. Acad. Sci. U. S. A. 2005, 102(6), 1859–1864.

37.

Margueron

Justin

Ohno

Sharpe

M. L.

Son

Drury

W. J.

3rd Voigt

Martin

S. R.

Taylor

W. R.

De Marco

. Role of the Polycomb Protein EED in the Propagation of Repressive Histone Marks. Nature 2009, 461(7265), 762–767.

38.

Southall

S. M.

Wong

P. S.

Odho

Roe

S. M.

Wilson

J. R.

Structural Basis for the Requirement of Additional Factors for MLL1 Set Domain Activity and Recognition of Epigenetic Marks. Mol. Cell 2009, 33(2), 181–191.

39.

Odho

Southall

S. M.

Wilson

J. R.

Characterization of a Novel WDR5-Binding Site That Recruits RbBP5 through a Conserved Motif to Enhance Methylation of Histone H3 Lysine 4 by Mixed Lineage Leukemia Protein-1. J. Biol. Chem. 2010, 285(43), 32967–76.

40.

Patel

Vought

V. E.

Dharmarajan

Cosgrove

M. S.

A Conserved Arginine-Containing Motif Crucial for the Assembly and Enzymatic Activity of the Mixed Lineage Leukemia Protein-1 Core Complex. J. Biol. Chem. 2008, 283(47), 32162–32175.

41.

Dou

Milne

T. A.

Ruthenburg

A. J.

Lee

J. W.

Verdine

G. L.

Allis

C. D.

Roeder

R. G.

Regulation of MLL1 H3K4 Methyltransferase Activity by Its Core Components. Nat. Struct. Mol. Biol. 2006, 13(8), 713–719.

42.

Wysocka

Swigut

Milne

T. A.

Dou

Zhang

Burlingame

A. L.

Roeder

R. G.

Brivanlou

A. H.

Allis

C. D.

WDR5 Associates with Histone H3 Methylated at K4 and Is Essential for H3 K4 Methylation and Vertebrate Development. Cell 2005, 121(6), 859–872.

43.

Richon

V. M.

Johnston

Sneeringer

C. J.

Jin

Majer

C. R.

Elliston

Fred Jerva

Scott

M. P.

Copeland

R. A.

Chemogenetic Analysis of Human Protein Methyltransferases. Chem. Biol. Drug Des. 2011, 78(2), 199–210.

44.

Kelly

T. K.

De Carvalho

D. D.

Jones

P. A.

Epigenetic Modifications as Therapeutic Targets. Nat. Biotechnol. 2010, 28(10), 1069–1078.

45.

Vedadi

Arrowsmith

C. H.

Allali-Hassani

Senisterra

Wasney

G. A.

Biophysical Characterization of Recombinant Proteins: A Key to Higher Structural Genomics Success. J. Struct. Biol. 2010, 172(1), 107–119.

46.

Moulick

Rodina

Aguirre

Felts

Dingledine

Chiosis

High-Throughput Screening Fluorescence Polarization Assay for Tumor-Specific Hsp90. J. Biomol. Screen. 2007, 12(7), 915–924.

47.

Kawamoto

S. A.

Thompson

A. D.

Coleska

Nikolovska-Coleska

Wang

Analysis of the Interaction of BCL9 with Beta-Catenin and Development of Fluorescence Polarization and Surface Plasmon Resonance Binding Assays for This Interaction. Biochemistry 2009, 48(40), 9534–9541.

48.

Reindl

Graber

Strebhardt

Berg

Development of High-Throughput Assays Based on Fluorescence Polarization for Inhibitors of the Polo-Box Domains of Polo-Like Kinases 2 and 3. Anal. Biochem. 2009, 395(2), 189–194.

49.

Collazo

Couture

J. F.

Bulfer

Trievel

R. C.

A Coupled Fluorescent Assay for Histone Methyltransferases. Anal. Biochem. 2005, 342(1), 86–92.

50.

Vedadi

Barsyte-Lovejoy

Liu

Rival-Gervier

Allali-Hassani

Labrie

Wigle

T. J.

DiMaggio

P. A.

Wasney

G. A.

Siarheyeva

. A chemical probe selectively inhibits G9a and GLP methyltransferase activity in cells. Nat. Chem. Biol. 2011, 7(8):566-74.

51.

Liu

Chen

Allali-Hassani

Quinn

A. M.

Wasney

G. A.

Dong

Barsyte

Kozieradzki

Senisterra

Chau

. Discovery of a 2,4-Diamino-7-Aminoalkoxyquinazoline as a Potent and Selective Inhibitor of Histone Lysine Methyltransferase G9a. J. Med. Chem. 2009, 52(24), 7950–7953.

52.

Liu

Chen

Allali-Hassani

Quinn

A. M.

Wigle

T. J.

Wasney

G. A.

Dong

Senisterra

Chau

Siarheyeva

. Protein Lysine Methyltransferase G9a Inhibitors: Design, Synthesis, and Structure Activity Relationships of 2,4-Diamino-7-Aminoalkoxy-Quinazolines. J. Med. Chem. 2010, 53(15), 5844–5857.

53.

Palmer

N. A.

Sattler

S. E.

Saathoff

A. J.

Sarath

A Continuous, Quantitative Fluorescent Assay for Plant Caffeic Acid O-Methyltransferases. J. Agric. Food Chem. 2010, 58(9), 5220–5226.

54.

Dorgan

K. M.

Wooderchak

W. L.

Wynn

D. P.

Karschner

E. L.

Alfaro

J. F.

Cui

Zhou

Z. S.

Hevel

J. M.

An Enzyme-Coupled Continuous Spectrophotometric Assay for S-Adenosylmethionine-Dependent Methyltransferases. Anal. Biochem. 2006, 350(2), 249–255.

55.

Hendricks

C. L.

Ross

J. R.

Pichersky

Noel

J. P.

Zhou

Z. S.

An Enzyme-Coupled Colorimetric Assay for S-Adenosylmethionine-Dependent Methyltransferases. Anal. Biochem. 2004, 326(1), 100–105.

56.

Kubicek

O’Sullivan

R. J.

August

E. M.

Hickey

E. R.

Zhang

Teodoro

M. L.

Rea

Mechtler

Kowalski

J. A.

Homon

C. A.

. Reversal of H3K9me2 by a Small-Molecule Inhibitor for the G9a Histone Methyltransferase. Mol. Cell 2007, 25(3), 473–481.

57.

Quinn

A. M.

Allali-Hassani

Vedadi

Simeonov

A Chemiluminescence-Based Method for Identification of Histone Lysine Methyltransferase Inhibitors. Mol. Biosyst. 2010, 6(5), 782–788.

58.

Cheng

Yadav

King

R. W.

Swanson

M. S.

Weinstein

E. J.

Bedford

M. T.

Small Molecule Regulators of Protein Arginine Methyltransferases. J. Biol. Chem. 2004, 279(23), 23892–23899.

59.

Wigle

T. J.

Provencher

L. M.

Norris

J. L.

Jin

Brown

P. J.

Frye

S. V.

Janzen

W. P.

Accessing Protein Methyltransferase and Demethylase Enzymology Using Microfluidic Capillary Electrophoresis. Chem. Biol. 2010, 17(7), 695–704.

60.

Gowher

Zhang

Cheng

Jeltsch

Avidin Plate Assay System for Enzymatic Characterization of a Histone Lysine Methyltransferase. Anal. Biochem. 2005, 342(2), 287–291.

61.

Dhayalan

Dimitrova

Rathert

Jeltsch

A Continuous Protein Methyltransferase (G9a) Assay for Enzyme Activity Measurement and Inhibitor Screening. J. Biomol. Screen. 2009, 14(9), 1129–1133.

62.

Min

Lunin

V. V.

Antoshenko

Dombrovski

Zeng

Allali-Hassani

Campagna-Slater

Vedadi

Arrowsmith

C. H.

. Structural Biology of Human H3K9 Methyltransferases. PLoS One 2010, 5(1), e8570.

63.

Zhang

J. H.

Chung

T. D.

Oldenburg

K. R.

A Simple Statistical Parameter for Use in Evaluation and Validation of High Throughput Screening Assays. J. Biomol. Screen. 1999, 4(2), 67–73.

64.

Nimura

Ura

Kaneda

Histone Methyltransferases: Regulation of Transcription and Contribution to Human Disease. J. Mol. Med. 2010, 88(12), 1213–1220.

65.

Avdic

Zhang

Lanouette

Groulx

Tremblay

Brunzelle

Couture

J. F.

Structural and Biochemical Insights into MLL1 Core Complex Assembly. Structure 2011, 19(1), 101–108.

66.

Crea

Hurt

E. M.

Mathews

L. A.

Cabarcas

S. M.

Sun

Marquez

V. E.

Danesi

Farrar

W. L.

Pharmacologic Disruption of Polycomb Repressive Complex 2 Inhibits Tumorigenicity and Tumor Progression in Prostate Cancer. Mol. Cancer 2011, 10(1), 40.

67.

Bian

Yang

Galka

Ouyang

Chen

Qiu

Liu

Jones

A. E.

MacKenzie

. Binding of Different Histone Marks Differentially Regulates the Activity and Specificity of Polycomb Repressive Complex 2 (PRC2). Proc. Natl. Acad. Sci. U. S. A. 2010, 107(45), 19266–19271.

68.

Patnaik

Chin

H. G.

Esteve

P. O.

Benner

Jacobsen

S. E.

Pradhan

Substrate Specificity and Kinetic Mechanism of Mammalian G9a Histone H3 Methyltransferase. J. Biol. Chem. 2004, 279(51), 53248–53258.

69.

Xiao

Jing

Wilson

J. R.

Walker

P. A.

Vasisht

Kelly

Howell

Taylor

I. A.

Blackburn

G. M.

Gamblin

S. J.

Structure and Catalytic Mechanism of the Human Histone Methyltransferase SET7/9. Nature 2003, 421(6923), 652–656.

70.

Couture

J. F.

Collazo

Hauk

Trievel

R. C.

Structural Basis for the Methylation Site Specificity of SET7/9. Nat. Struct. Mol. Biol. 2006, 13(2), 140–146.

71.

Trievel

R. C.

Beach

B. M.

Dirk

L. M.

Houtz

R. L.

Hurley

JH.

Structure and Catalytic Mechanism of a Set Domain Protein Methyltransferase. Cell 2002, 111(1), 91–103.

72.

Dirk

L. M.

Flynn

E. M.

Dietzel

Couture

J. F.

Trievel

R. C.

Houtz

R. L.

Kinetic Manifestation of Processivity during Multiple Methylations Catalyzed by Set Domain Protein Methyltransferases. Biochemistry 2007, 46(12), 3905–3915.

73.

Couture

J. F.

Collazo

Brunzelle

J. S.

Trievel

R. C.

Structural and Functional Analysis of SET8, a Histone H4 Lys-20 Methyltransferase. Genes Dev. 2005, 19(12), 1455–1465.

74.

Pahlich

Bschir

Chiavi

Belyanskaya

Gehring

Different Methylation Characteristics of Protein Arginine Methyltransferase 1 and 3 toward the Ewing Sarcoma Protein and a Peptide. Proteins 2005, 61(1), 164–175.

Supplementary Material

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Fluorescence-Based Methods for Screening Writers and Readers of Histone Methyl Marks

Abstract

Keywords

Introduction

Material and Methods

SAHH Assay

HMT Assay

Z Factor Determination

Fluorescence Polarization

Results and Discussion

Assay and Screening Writers of Histone Methyl Markers

Assay Optimization

Determining the Kinetic Parameters for HMTs

HTS Using Continuous Assay

Screening Histone Methyl Marks Using Peptide Displacement

Footnotes

Acknowledgements

References

Supplementary Material