This work has been supported by the DFG grant JE 252/7.

1. Preparation of Peptide Arrays
<ol>
	<li>Programming of the Peptide Sequences
	<ol>
		<li>Design peptide sequences for the synthesis using the MultiPep Spotter software. Enter the peptide sequences in single letter codes. 
		Peptide 1: T A R K S T G G K A</li>
		<li>Generate alanine or arginine walk libraries with a specific command (.replace, A) to screen the important amino acids required for the recognition of a defined substrate. For instance in the following example the first line is the wild type sequence and from the second line each amino acid in a peptide is altered to alanine sequentially. 
		replace, A 
		T-A-R-K-S-T-G 
		A-A-R-K-S-T-G 
		T-A-R-K-S-T-G 
		T-A-A-K-S-T-G 
		T-A-R-A-S-T-G 
		T-A-R-K-A-T-G 
		T-A-R-K-S-A-G 
		T-A-R-K-S-T-A</li>
		<li>Use similar commands (.replace R, .replace N etc.) as described in 1.1.2 with all the naturally available amino acid residues (eventually omitting cysteine, methionine and tryptophan, because these residues are prone to oxidation and sometimes lower synthesis yield) to&#160;synthesize a peptide array for determining the consensus substrate sequence motif for an enzyme. 
		NOTE: As shown in Figure 4A, the horizontal axis represents the given peptide sequence and the vertical axis represents the amino acids that are selected for command. Similar to the sequence shown in 1.1.2, A on the vertical axis means that an Ala sequentially replaces&#160;&#160;each position of the given sequence in the first row in Fig. 4A. Similarly arginine sequentially replaces each amino acid in the second&#160;row and so on with the other amino acids. Complete peptide array libraries are synthesized by systematically replacing each residue in the provided peptide substrate sequence with each of the other naturally available amino acid residues, this helps to understand the influence of each residue at each position of the given sequence.</li>
		<li>Alternatively, methylation of known or predicted peptide substrates can be easily verified synthesizing peptide libraries with the target peptides of the PKMT together with positive and negative methylation controls (i.e. peptides known to methylated or known not to be methylated). Synthesize peptides by exchanging the putative target lysine to alanine to confirm methylation of the target lysine as shown below by manual programming. 
		Wild type peptide: S T G G K P R Q F L 
		Mutant peptide : S T G G A P R Q F L 
		NOTE: The software also has inherent scripts to create various libraries, like epitope mapping with a desired frame shift of the sequence to map important amino acids required for an interaction.&#160;</li>
	</ol>
	</li>
	<li>Preparation of Membrane, Amino Acids and Reagents
	<ol>
		<li>Pre-swell the SPOT membrane in DMF (N,N-Dimethylformamide) for 10 min and then place the wet membrane on the SPOT synthesizer frame without air bubbles or wrinkles.</li>
		<li>Wash the membrane three times with 100% ethanol. Dry the membranes completely for at least 10 min before starting the synthesis program.</li>
		<li>In parallel, freshly prepare 0.5 M working concentrations of commercially available Fmoc-protected amino acids by dissolving them in NMP and also prepare the activator and base as described in Table 1.</li>
		<li>Make sure that all the waste solution containers of the SPOT synthesizer were emptied and refill the solvent reservoirs with the DMF, ethanol and piperidine. 
		NOTE: Now the machine is ready for the synthesis.</li>
	</ol>
	</li>
	<li>Spotting of the First Amino Acid
	<ol>
		<li>To generate the amide bond with the free amino group on the membrane, activate the carboxy group of the incoming amino acid by adding the activator (N,N&#8217;-Diisopropylcarbodiimide) and base solution (Ethyl hydroxyimino cyanoacetate) to the amino acid derivative.</li>
		<li>Spot the desired volumes of activated amino acids on the Amino-PEG functionalized membrane using the programmable robot. 
		NOTE: Thereby, the C-terminus of the amino acid is coupled to the amino group of the membrane.</li>
		<li>Repeat this step three times to ensure better coupling of the first activated amino acid. Incubate the membrane for 20 min and then wash with DMF to remove uncoupled amino acids.</li>
	</ol>
	</li>
	<li>Blocking of Free Spaces on Non-spot Areas 
	NOTE: After the coupling of the first C-terminal amino acid of all peptides to the amino group of the membrane, the amino groups between the spots and also some of the amino groups within the spot areas do not form bonds with the amino acids.
	<ol>
		<li>Block these free amino groups by incubating with the capping solution (20% acetic anhydride in DMF) for 5 min. 
		NOTE: This avoids the coupling of amino acids in the further cycles with the membrane instead of the growing peptide chain.</li>
	</ol>
	</li>
	<li>Removal of Fmoc Group
	<ol>
		<li>After the blocking, wash the membrane with DMF 4 times and then incubate with 20% piperidine in DMF for 20 min to remove the Fmoc protection group. 
		NOTE: This step leads to removal of the amino-protecting Fmoc groups and it is called deprotection.</li>
		<li>Afterwards, wash the membrane twice with DMF and twice with ethanol and allow it to dry. 
		NOTE: Now the membrane is ready for the coupling of the next amino acids.&#160;</li>
	</ol>
	</li>
	<li>Chain Elongation 
	NOTE: Addition of each amino acid is called a cycle. Apart from the coupling of first amino acid, the cycle begins with the deprotection of Fmoc group of the coupled amino acid and it is followed by the coupling of the activated carboxyl group of an incoming amino acid to the amino group of the growing peptide chain.
	<ol>
		<li>Repeat steps 1.3 and 1.5 until the desired peptide length is achieved.</li>
	</ol>
	</li>
	<li>Staining 
	NOTE: After the final cycle, the peptide arrays are stained with bromophenol blue to confirm the successful synthesis of peptides. Bromophenol blue binds with the free amino groups of the peptides. Peptide spots show light blue color after the staining but the intensity of the spots varies depending on the peptide sequence. This staining allows confirmation of the complete removal of piperidine, because in presence of piperidine bromophenol blue does not stain the peptides. In addition, this also helps to mark the peptide arrays for further usage.
	<ol>
		<li>After the Fmoc group deprotection in the last synthesis cycle, wash the membrane with DMF and 100% ethanol. Then, treat the membrane with bromophenol blue (0.02% bromophenol blue in DMF) for a minimum of 5 min until it completely turns blue.</li>
		<li>Afterwards, wash the membrane with DMF and ethanol followed by drying for 10 min. Now take out the membrane from the synthesizer and perform side chain deprotection (section 1.8) manually. If needed, photograph the membrane to document the results of the peptide synthesis (Fig. 2).&#160;</li>
	</ol>
	</li>
	<li>Side Chain Deprotection
	<ol>
		<li>Treat the membranes with 20 to 25 ml of side chain deprotection mixture consisting of 95% Trifluoroacetic acid (TFA) to cleave the side chain protection groups and scavenger reagents (2.5% water and 2.5% triisopropylsilane) to protect the side chains of amino acids from modification during this step. Take a sufficient volume of deprotection mixture to ensure that it covers the membrane completely in a tightly closed chemical resistant box and incubate it for 1 to 2 hr while shaking gently.</li>
		<li>Wash the membrane six times with 20-25 ml of DCM (Dichloromethane) for 2 min each and finally twice with 100% ethanol and dry it in a desiccator overnight. 
		NOTE: Now the membrane can be used for experiments. A scheme of the peptide array synthesis is provided in Figure 1.&#160;</li>
	</ol>
	</li>
</ol>
2. Protein Expression and Purification
<ol>
	<li>Protein Expression of PKMTs as GST Fusions
	<ol>
		<li>Using standard techniques, clone the gene encoding the full length PKMT or its catalytic domain into a bacterial expression vector (like pGEX-6p2) to express it as GST-fusion protein.</li>
		<li>Transfer the vector with the desired PKMT insert into BL21 or BL21 codon plus E. coli cells by the heat shock method or any other method.</li>
		<li>Prepare a pre-culture with 30 ml of Luria-Bertani (LB) media on the day of expression and incubate at 37 &#176;C for 7 to 8 hr with continuous shaking.</li>
		<li>Next, transfer 10 ml of the pre-culture into a 2 L big baffled flasks containing 1 L of LB media and incubate at 37 &#176;C in the incubator with continuous shaking until the culture reaches a defined optical density at 600 nm. 
		NOTE: While this can be optimized for each protein, we routinely induce at an OD600nm of about 0.8. The induction temperature has to be optimized for each protein individually, some proteins show good expression at 22 &#176;C and some at higher temperatures.</li>
		<li>Shift the cells to the induction temperature for 15 min, then induce with 1 mM of isopropyl-beta-D-thiogalactopyranoside and allow the culture to grow for 10-12 hr at induction temperature.</li>
		<li>Afterwards harvest the cells by centrifugation at 5,000 x g for 15 min and finally wash the pellet with 30 ml of STE buffer (10 mM Tris pH 8.0, 100 mM NaCl and 0.1 mM EDTA) and store at -20 &#176;C for further usage.</li>
	</ol>
	</li>
	<li>Protein Purification&#160;
	<ol>
		<li>Thaw the cell pellet and re-suspend in 30 ml of sonication buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1 mM DTT and 5% glycerol) and disrupt by sonication. 
		NOTE: This buffer needs to be optimized for each protein eventually.</li>
		<li>Centrifuge the cell lysate at 22,000 x g for 1 hr and, later collect the supernatant and pass through a column containing 600 &#181;l of glutathione Sepharose 4B resin.</li>
		<li>Wash the column with 50 ml of sonication buffer and next with 100 ml of high salt buffer (50 mM Tris pH 7.5, 500 mM NaCl, 1 mM DTT and 5% glycerol) to remove unspecifically bound proteins. Elute the bound protein with 5 ml of high salt buffer containing 40 mM glutathione.</li>
		<li>Dialyze the eluted protein in 2 L of dialysis buffer with low glycerol (20 mM Tris pH 7.4, 100 mM KCl, 0.5 mM DTT and 10% glycerol) for 2 hr and later in 20 mM Tris pH 7.4, 100 mM KCl, 0.5 mM DTT and 70% glycerol for 8 hr or overnight. Make sure all the purification buffers are at 4 &#176;C and perform the protein purification in the cold room to avoid protein denaturation.</li>
	</ol>
	</li>
</ol>
3. Peptide Array Methylation
<ol>
	<li>Pre-incubation of Peptide Arrays
	<ol>
		<li>Perform peptide array methylation either in a box of appropriate size or in a plastic bag to avoid the wastage of enzyme and expensive radioactively labelled AdoMet. Pre-incubate the peptide array membrane in sealed plastic bag containing the respective methylation buffer without enzyme and labeled [methyl-3H]-AdoMet for 10 min. The volume of buffer depends upon the size of the array, for example use 8 ml of methylation buffer for a full specificity profile peptide array. Optimize the amount and concentration of the AdoMet for each enzyme.</li>
	</ol>
	</li>
	<li>Methylation of Peptide Arrays
	<ol>
		<li>Discard the pre-incubation buffer and incubate the membrane in 8 ml of methylation buffer containing the respective PKMT and labeled [methyl-3H]-AdoMet for 1 to 2 hr. 
		NOTE: The amount of enzyme required for a methylation reaction depends upon the activity of specific PKMT, we initiate our screening experiments typically with 50 nM enzyme final concentration.</li>
	</ol>
	</li>
	<li>Washing of Peptide Arrays
	<ol>
		<li>Afterwards discard the methylation buffer in a radioactive waste container and wash the peptide arrays for 5 times with 20 ml of buffer containing 100 mM ammonium bicarbonate and 1% SDS for 5 min to remove the bound protein. Discard the washing buffer in a radioactive waste container.</li>
	</ol>
	</li>
	<li>Detection of the Radioactive Signal
	<ol>
		<li>After the washes incubate the membrane for 5 min&#160;in 10 ml of amplify solution (NAMP100V).</li>
		<li>Then, discard the amplify solution into the radioactive waste container for organic solvents, seal the peptide array in a plastic bag and put it in an autoradiography cassette.</li>
		<li>Place the autoradiography film on the peptide array in the dark room. Close the cassette carefully and expose it at -80 &#176;C for the necessary time. Then, develop the film to analyze the results. Capture the image couple of times with different exposition times to avoid saturation of strongly methylated peptide substrates.</li>
	</ol>
	</li>
</ol>
4. Data Processing and Analysis
<ol>
	<li>Densitometric Analysis of Spot Intensities
	<ol>
		<li>Scan the film in a conventional scanner and analyze the spot intensities by densitometry. Do this using ImageJ (which is freeware) or a commercial program. 
		NOTE: We use Phoretix software for this task.</li>
	</ol>
	</li>
	<li>Normalization and Averaging of Results from Different Membranes
	<ol>
		<li>Conduct the peptide array methylation experiments at least in triplicate. Normalize the scanned intensities for each experiment by background subtraction and setting the activity at a reference substrate as 1.0. Average the data for the individual spots and report them as mean values and standard deviations.</li>
	</ol>
	</li>
	<li>Data Analysis and Display
	<ol>
		<li>Plot the data in 2D using a grayscale or color scheme to indicate the relative activity. Do this with Excel or Sigmaplot as shown here. Also, prepare a plot of the distribution of the standard deviations of the repeated experiments to analyze the quality of the data.</li>
		<li>To compare and display the accuracy of the recognition of each residue in the substrate peptide quantitatively, calculate the relative preference of the PKMT for each amino acid i at position x by a discrimination factor D 16,17: 
		DX;i = &#60; Vj&#8800;i&#62;/Vi - 1 
		Where vi is the rate of methylation of the peptide variant carrying amino acid i at position x and &#60;vj&#8800;i&#62; is the average rate of methylation of all 19 peptides carrying a different amino acid j&#8800;i at position x (including the wild type sequence). 
		NOTE: The discrimination factor defined by this depends on the background level. If only one residue is accepted at a certain position with a background of 3%, a discrimination factor of 32.3 is obtained. No preference at a position results in a discrimination factor of 0.</li>
	</ol>
	</li>
</ol>

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No conflicts of interest declared.

SPOT synthesis as described here is a powerful method to map protein-protein interaction sites and investigate the substrate recognition of peptide modifying enzymes. However, SPOT synthesis still has certain drawbacks, because although the peptides synthesized by the SPOT method are reported to have more than 90% purity 21 this is difficult to confirm in each instance. Therefore, results have to be reproduced by other methods for instance using protein domains or with purified peptides synthesized by standard...

In the last two decades, several reports demonstrated the importance of post-translational modifications (PTM) in cellular development and several diseases like cancer, but recently protein lysine methylation has emerged as an another vital PTM. While initially histone lysine methylation was found to be an essential chromatin mark, later work also showed lysine methylation of several non-histone proteins 1-4. The sequential transfer of methyl groups from S-adenosyl-L-methionine to the &#949;-amino group of lysine residues is catalyzed by a family of enzymes called Protein Lysine Methyltransferases (PKMTs) that contains over 60 proteins in the human genome. PKMTs were initially discovered as histone modifying enzymes that methylate specific lysine residue but later reports demonstrated that they could also methylate non-histone proteins 5. Up to now, approximately 5,000 lysine methylation sites were identified on different proteins 6, but the enzymes responsible for these modifications are often not identified. One reason for this is that the specificity of most PKMTs has not been studied extensively. Therefore, it recurrently occurs that novel substrates of PKMTs are discovered. The lack of a detailed knowledge of the substrate specificity of PKMTs hinders understanding of their biological function and role. To study the specificity of a PKMT in detail, the methylation rates of many peptide substrates that differ in one or few amino acids must be measured and compared, which is ideally done using peptide arrays. Based on the resulting specificity profiles, potential substrates of PKMTs can be identified that can be studied further.
Peptide arrays are widely used tools for the biochemical analysis of antibodies, peptide modifying enzymes and mapping of protein-protein interaction sites (antibody-antigen, receptor-ligand) 7-9. Several hundreds of peptides are needed for such applications. Different methods are available for peptide synthesis, among them peptide synthesis on resin is very commonly used, but it has limitations in throughput and it is relatively expensive. These issues were resolved with the introduction of the SPOT synthesis method by Frank and colleagues 10. The SPOT synthesis method allows synthesis of several hundred peptides in parallel and on average it is inexpensive compared to resin synthesis. The peptides synthesized on cellulose membrane can be used either directly for various applications or peptides can be cleaved from the membrane and used as free peptides for in solution assays or to prepare peptide microarrays 10-13.
SPOT synthesis is a variant of the solid phase peptide synthesis, which uses a cellulose membrane as a solid support and employs the standard Fmoc-chemistry 10-13. Hence, the synthesis of peptide chains starts at the C-terminal end and proceeds towards the N-terminal end in contrast to the biological synthesis in ribosomes. Cellulose membranes are functionalized for the attachment of the first activated amino acids (Fig. 1). The SPOT method is based on the sequential delivery of activated amino acids in a droplet of solvent to defined spots on the membrane using an automated pipetting system. The droplet of liquid is dispensed on the porous membrane where it forms a circular wet spot, which later acts as an open reactor for the chemical reactions in peptide synthesis. The spot size is determined by the volume dispensed and the absorptive capacity of the membrane, multiples of such spots are arranged as arrays. The scale of synthesis correlates with the spot size and the loading capacity of the membrane. The distance between the spots and the density of arrays are managed by varying the spot sizes. Cellulose membrane has several advantages as solid phase in peptide synthesis, it is inexpensive, tolerant to the chemicals used in peptide synthesis, stable in aqueous solutions and easy to handle. In addition, its hydrophilic nature makes it suitable for several biological assay systems. SPOT synthesis can be carried out manually or automated (for 1000s of peptides) depending on the required number of peptides. A fully automated SPOT synthesizer from Intavis (K&#246;ln, Germany) is used for our applications. It permits synthesis of peptides in different amounts and of different length. Linear peptides are regularly synthesized with 15 to 20 amino acid length, in addition peptides of up to 42 amino acid can also be prepared by step-wise synthesis 14,15. However, increasing the number of amino acids leads to reductions in the overall coupling yields, which affects the quality of the peptides. Because of the low amount of peptides per spot, the products are often difficult to purify and the quality of individual peptides cannot be easily assessed. Therefore, the results obtained from SPOT peptide arrays must be confirmed either with peptides synthesized by standard methods in larger scale, which can be purified and analyzed according to standards in peptide synthesis or by synthesizing the proteins containing the desired peptide sequences. Still, we found the SPOT synthesis to be highly reliable and results generally to be reproducible. SPOT synthesis is not restricted to proteinogenic amino acids, several commercially available modified amino acids also can be used for synthesis, allowing peptides to be modified before and after the final cleavage of the side-chain protection group and, furthermore, it also allows incorporation of phosphorylated, methylated or acetylated amino acids 11.
Immobilized peptide libraries synthesized by the SPOT method can be directly used for many biological and biochemical assays. We employed peptide arrays comprising 300-400 peptides to investigate the substrate specificity of PKMTs. For enzymatic modification, the peptide arrays are incubated with the respective PKMT and labeled [methyl-3H]-AdoMet in an appropriate buffer. The methylation of the respective substrate is analyzed by following the enzymatic transfer of the radioactively labelled methyl groups from AdoMet to the peptide substrate via autoradiography (Fig. 3). By this procedure, the peptide arrays allow the study of methylation of different peptide substrates at the same time. One important advantage of this method is that all the peptides are methylated in competition, such that during the linear phase of the methylation kinetics, the relative methylation of each peptide is proportional to the catalytic rate constant divided by the dissociation constant (kcat/KD) of the enzyme for the respective peptide substrate. Therefore, the amount of radioactivity incorporated into each spot is directly correlated with the enzymatic activity towards the particular peptide. Using the results of a peptide array methylation experiment, the specificity profile of the PKMT can be defined and based on this novel substrates can be predicated. Peptide arrays allow the rapid and cost efficient validation of the methylation of novel substrates at the peptide level. For this, arrays are prepared that contain the predicted novel substrates together with modified peptides containing an Ala instead of Lys at the target sites as well as positive and negative control peptides. Finally, the novel substrates can be prepared as proteins together with mutants, in which the target Lys is altered to Ala and the methylation can be confirmed at the protein level. Depending on the results, this is then followed by biological studies addressing potential roles of the methylation of the newly described protein substrates.

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			<td height="21" style="height:21px;width:325px;">Name of Material/ Equipment</td>
			<td style="width:112px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:179px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ethanol abs Gradient Grade HPLC</td>
			<td>Honeywell</td>
			<td>10299901</td>
			<td>Flammable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N,N-Dimethylformamide Peptide Synthesis</td>
			<td>Biosolve</td>
			<td style="width:119px;">4193302</td>
			<td>Flammable, Toxic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Piperidine &#8805; 99 % for Peptide Synthesis</td>
			<td>Roth</td>
			<td>A122.1</td>
			<td>Flammable, Toxic, corrosive</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Oxyma Pure (Ethyl (hydroxyimino)cyanoacetate )</td>
			<td>Novabiochem</td>
			<td>8510860100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">N,N&#8217;-Diisopropylcarbodiimide purum &#8805; 98% GC</td>
			<td>Fluka</td>
			<td>38370</td>
			<td>Flammable, Toxic, corrosive</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Acetic anhydride</td>
			<td>Roth</td>
			<td>CP28.1</td>
			<td>Flammable, Toxic, corrosive</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triisopropylsilane 99%,</td>
			<td>Aldrich</td>
			<td>233781</td>
			<td>Flammable, Toxic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dichlormethane &#8805;99,9%</td>
			<td>Roth</td>
			<td>P089.1</td>
			<td>Carcinogenic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N-Methyl-2-Pyrrolidone</td>
			<td>Roth</td>
			<td>4306.2</td>
			<td>Toxic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Derivatized cellulose Membrane</td>
			<td style="width:112px;">Intavis AG K&#246;ln</td>
			<td style="width:119px;">32.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Trifluoroacetic acid</td>
			<td>Roth</td>
			<td>P088.2</td>
			<td>Toxic, corrosive</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Bromphenolblue</td>
			<td>AppliChem</td>
			<td>A3640.0010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ammonium Hydrogen Carbonate</td>
			<td>Roth</td>
			<td>T871.2</td>
			<td>Toxic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Dodecyl Sulfate Pellets</td>
			<td>Roth</td>
			<td>CN30.3</td>
			<td>Toxic, Flammable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">MultiPep Synthesizer</td>
			<td style="width:112px;">Intavis AG K&#246;ln</td>
			<td style="width:119px;">n.a.</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">HyperfilmTM high performance film</td>
			<td style="width:112px;">GE Healthcare</td>
			<td style="width:119px;">28906837</td>
			<td style="width:179px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:325px;">Phoretix software</td>
			<td style="width:112px;">TotalLab</td>
			<td style="width:119px;">n.a.</td>
			<td style="width:179px;"></td>
		</tr>
	</tbody>
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specificity analysis of protein lysine methyltransferases using spot peptide arrays

Lysine methylation is an emerging post-translation modification and it has been identified on several histone and non-histone proteins, where it plays crucial roles in cell development and many diseases. Approximately 5,000 lysine methylation sites were identified on different proteins, which are set by few dozens of protein lysine methyltransferases. This suggests that each PKMT methylates multiple proteins, however till now only one or two substrates have been identified for several of these enzymes. To approach this problem, we have introduced peptide array based substrate specificity analyses of PKMTs. Peptide arrays are powerful tools to characterize the specificity of PKMTs because methylation of several substrates with different sequences can be tested on one array. We synthesized peptide arrays on cellulose membrane using an Intavis SPOT synthesizer and analyzed the specificity of various PKMTs. Based on the results, for several of these enzymes, novel substrates could be identified. For example, for NSD1 by employing peptide arrays, we showed that it methylates K44 of H4 instead of the reported H4K20 and in addition H1.5K168 is the highly preferred substrate over the previously known H3K36. Hence, peptide arrays are powerful tools to biochemically characterize the PKMTs.

Peptide arrays were successfully used to biochemically characterize the specificity of PKMTs, and several novel histone and non-histone substrates of PKMT were identified by this approach 5,16-19. Defining the correct substrate spectrum of a PKMT (or any enzyme) is an essential step towards understanding of its molecular mechanism and cellular functions.
As an example of the application of the peptide SPOT array methylation method, we describe the results of the specificity analysis...

Watch this Scientific Journal Video about Specificity Analysis of Protein Lysine Methyltransferases Using SPOT Peptide Arrays at JoVE.com

Specificity Analysis of Protein Lysine Methyltransferases Using SPOT Peptide Arrays

Peptide arrays synthesized by the SPOT method can be used to analyze the substrate specificity of Protein lysine methyltransferases (PKMTs) and to define the substrate spectrum of PKMTs to understand their biological role. This protocol describes how to synthesize peptide arrays, methylate them with PKMTs, and analyze the results.

Lysine methylation is an emerging post-translation modification and it has been identified on several histone and non-histone proteins, where it plays ...

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Research

JoVE Journal

Biology

13.8K Views.  Stuttgart University. Peptide arrays synthesized by the SPOT method can be used to analyze the substrate specificity of Protein lysine methyltransferases (PKMTs) and to define the substrate spectrum of PKMTs to understand their biological role. This protocol describes how to synthesize peptide arrays, methylate them with PKMTs, and analyze the results.

Video: Specificity Analysis of Protein Lysine Methyltransferases Using SPOT Peptide Arrays

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Quantification of Proteins Using Peptide Immunoaffinity Enrichment Coupled with Mass Spectrometry

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in quantitation, are associated with a high cost, long lead time, and are fraught with drawbacks (e.g. heterophilic antibodies, autoantibody interference, 'hook-effect').1 An alternative technique is affinity enrichment of peptides coupled with quantitative mass spectrometry, commonly referred to as SISCAPA (Stable Isotope Standards and Capture by Anti-Peptide Antibodies).2 In this technique, affinity enrichment of peptides with stable isotope dilution and detection by selected/multiple reaction monitoring mass spectrometry (SRM/MRM-MS) provides quantitative measurement of peptides as surrogates for their respective proteins. SRM/MRM-MS is well established for accurate quantitation of small molecules 3, 4 and more recently has been adapted to measure the concentrations of proteins in plasma and cell lysates.5-7 To achieve quantitation of proteins, these larger molecules are digested to component peptides using an enzyme such as trypsin. One or more selected peptides whose sequence is unique to the target protein in that species (i.e. "proteotypic" peptides) are then enriched from the sample using anti-peptide antibodies and measured as quantitative stoichiometric surrogates for protein concentration in the sample. Hence, coupled to stable isotope dilution (SID) methods (i.e. a spiked-in stable isotope labeled peptide standard), SRM/MRM can be used to measure concentrations of proteotypic peptides as surrogates for quantification of proteins in complex biological matrices. The assays have several advantages compared to traditional immunoassays. The reagents are relatively less expensive to generate, the specificity for the analyte is excellent, the assays can be highly multiplexed, enrichment can be performed from neat plasma (no depletion required), and the technique is amenable to a wide array of proteins or modifications of interest.8-13 In this video we demonstrate the basic protocol as adapted to a magnetic bead platform.

Stable Isotope Standards and Capture by Anti-Peptide Antibodies (SISCAPA) couples affinity enrichment of peptides with stable isotope dilution mass spectrometry (MRM-MS) to provide quantitative measurement of peptides as surrogates for their respective proteins. Here we describe the protocol using magnetic particles in a partially automated format.

There is a great need for quantitative assays in measuring proteins. Traditional sandwich immunoassays, largely considered the gold standard in ...

Application of MassSQUIRM for Quantitative Measurements of Lysine Demethylase Activity

Recently, epigenetic regulators have been discovered as key players in many different diseases 1-3. As a result, these enzymes are prime targets for small molecule studies and drug development 4. Many epigenetic regulators have only recently been discovered and are still in the process of being classified. Among these enzymes are lysine demethylases which remove methyl groups from lysines on histones and other proteins. Due to the novel nature of this class of enzymes, few assays have been developed to study their activity. This has been a road block to both the classification and high throughput study of histone demethylases. Currently, very few demethylase assays exist. Those that do exist tend to be qualitative in nature and cannot simultaneously discern between the different lysine methylation states (un-, mono-, di- and tri-). Mass spectrometry is commonly used to determine demethylase activity but current mass spectrometric assays do not address whether differentially methylated peptides ionize differently. Differential ionization of methylated peptides makes comparing methylation states difficult and certainly not quantitative (Figure 1A). Thus available assays are not optimized for the comprehensive analysis of demethylase activity.
Here we describe a method called MassSQUIRM (mass spectrometric quantitation using isotopic reductive methylation) that is based on reductive methylation of amine groups with deuterated formaldehyde to force all lysines to be di-methylated, thus making them essentially the same chemical species and therefore ionize the same (Figure 1B). The only chemical difference following the reductive methylation is hydrogen and deuterium, which does not affect MALDI ionization efficiencies. The MassSQUIRM assay is specific for demethylase reaction products with un-, mono- or di-methylated lysines. The assay is also applicable to lysine methyltransferases giving the same reaction products. Here, we use a combination of reductive methylation chemistry and MALDI mass spectrometry to measure the activity of LSD1, a lysine demethylase capable of removing di- and mono-methyl groups, on a synthetic peptide substrate 5. This assay is simple and easily amenable to any lab with access to a MALDI mass spectrometer in lab or through a proteomics facility. The assay has ~8-fold dynamic range and is readily scalable to plate format 5.

We present a method for using MALDI mass spectrometry and reductive methylation chemistry to quantify changes in lysine methylation.

Recently, epigenetic regulators have been discovered as key players in many different diseases 1-3. As a result, these enzymes are prime targets for small ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

A High Throughput MHC II Binding Assay for Quantitative Analysis of Peptide Epitopes

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design1,2. Here, a peptide-MHC II binding assay is scaled to 384-well format. The scaled down protocol reduces reagent costs by 75% and is higher throughput than previously described 96-well protocols1,3-5. Specifically, the experimental design permits robust and reproducible analysis of up to 15 peptides against one MHC II allele per 384-well ELISA plate. Using a single liquid handling robot, this method allows one researcher to analyze approximately ninety test peptides in triplicate over a range of eight concentrations and four MHC II allele types in less than 48 hr. Others working in the fields of protein deimmunization or vaccine design and development may find the protocol to be useful in facilitating their own work. In particular, the step-by-step instructions and the visual format of JoVE should allow other users to quickly and easily establish this methodology in their own labs.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design. &#160;Here, a peptide-MHC II binding assay scaled to 384-well plates is described. This cost effective format should prove useful in the fields of protein deimmunization and vaccine design and development.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or ...

Identifying Protein-protein Interaction Sites Using Peptide Arrays

Protein-protein interactions mediate most of the processes in the living cell and control homeostasis of the organism. Impaired protein interactions may result in disease, making protein interactions important drug targets. It is thus highly important to understand these interactions at the molecular level. Protein interactions are studied using a variety of techniques ranging from cellular and biochemical assays to quantitative biophysical assays, and these may be performed either with full-length proteins, with protein domains or with peptides. Peptides serve as excellent tools to study protein interactions since peptides can be easily synthesized and allow the focusing on specific interaction sites. Peptide arrays enable the identification of the interaction sites between two proteins as well as screening for peptides that bind the target protein for therapeutic purposes. They also allow high throughput SAR studies. For identification of binding sites, a typical peptide array usually contains partly overlapping 10-20 residues peptides derived from the full sequences of one or more partner proteins of the desired target protein. Screening the array for binding the target protein reveals the binding peptides, corresponding to the binding sites in the partner proteins, in an easy and fast method using only small amount of protein.
In this article we describe a protocol for screening peptide arrays for mapping the interaction sites between a target protein and its partners. The peptide array is designed based on the sequences of the partner proteins taking into account their secondary structures. The arrays used in this protocol were Celluspots arrays prepared by INTAVIS Bioanalytical Instruments. The array is blocked to prevent unspecific binding and then incubated with the studied protein. Detection using an antibody reveals the binding peptides corresponding to the specific interaction sites between the proteins.

Peptide array screening is a high throughput assay for identifying protein-protein interaction sites. This allows mapping multiple interactions of a target protein and can serve as a method for identifying sites for inhibitors that target a protein. Here we describe a protocol for screening and analyzing peptide arrays.

Protein-protein interactions mediate most of the processes in the living cell and control homeostasis of the organism. Impaired protein interactions may ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...