This work was supported by the National Institutes of Health grants 5R01GM115446 and 1R01GM130840.

Mice used in this protocol were euthanized according to NIH guidelines. The Animal Welfare Committee at University of Texas Health Science Center in Houston approved all animal work.
1. Preparation of cell lysates
<ol>
	<li>Prepare lysis buffer as described in Table 1. To 10 mL of PBS, add 0.1 g of n-dodecyl &#946;-D-maltoside detergent (DDM) and rotate to dissolve. Add 100 &#181;L of phosphatase inhibitor cocktail 2, ML211 (10 &#181;M), PMSF (10 mM) and protease inhibitor cocktail (1x) and chill the buffer on ice before use.</li>
	<li>Transfer required media containing cells from the incubator into 15 mL or 50 mL conical tubes and spin cells at 350 x g for 5 min and aspirate to get rid of any cell debris. 
	NOTE: We used 1 x 107 cells for each acyl-RAC reaction to be performed.</li>
	<li>Wash the pellet by resuspending it in 5 mL of PBS and spinning at 350 x g for 5 min. 
	NOTE: Perform step 1.3 quickly to avoid cell lysis due to extended incubation in PBS.</li>
	<li>Add 600 &#181;L of lysis buffer prepared in step 1.1 to the pellet and lyse it by shaking at 1500 rpm in a thermal shaker for 30 min at 4 &#176;C.</li>
	<li>Clear lysates by centrifugation at 20,000 x g at 4 &#730;C for 30 min to pellet detergent-insoluble materials. Collect cleared lysate in pre-cooled 1.5 mL microfuge tubes and keep them on ice.</li>
	<li>Perform a Bradford/BCA assay to estimate protein concentration. It is critical to ensure the same amount of protein across different samples prior to performing the experiment. We recommend using at least 500 &#181;g of protein per reaction. 
	NOTE: In the experiments described, we used cultured (Jurkat) cells and primary splenocytes from mice tissue. The lysis method described above can be adopted to other cell types as well. The average protein concentration obtained for the abovementioned cell types is approximately 500 &#181;g per 1 x 107 cells. Jurkat cells were maintained in RPMI-1640 medium modified to contain 2 mM L-glutamine, 10 mM HEPES, 1 mM sodium pyruvate, 4,500 mg/L glucose, and 1,500 mg/L sodium bicarbonate supplemented with 10% FBS at 37 &#176;C with 5% CO2. We used 1 x 107 Jurkat cells per reaction. Primary splenocytes were isolated from mouse spleen tissue as described26. Briefly, spleen tissue was macerated on ice, followed by lysis of erythrocytes in hypotonic solution and separation from the lymphocytes by centrifugation. We used 1 x 107 primary cells per reaction.</li>
</ol>
2. Acyl-RAC: Blocking of free thiol groups
NOTE: All subsequent steps can be performed at room temperature (RT).
<ol>
	<li>Transfer lysate into a fresh 1.5 mL microfuge tube and perform chloroform-methanol (CM) precipitation as described below.
	<ol>
		<li>Add methanol (MeOH) and chloroform (CHCl3) to the lysate at a final ratio of lysate: MeOH: CHCl3 of 2:2:1 and shake vigorously to create a homogeneous suspension.</li>
		<li>Spin at 10,000 x g for 5 min to form a pellet (&#8220;pancake&#8221;) at the interphase between aqueous and organic phases.</li>
		<li>Tilt the tube and aspirate as much solvent as possible using a needle or a gel loading tip.</li>
		<li>Air dry the protein pellet for a few minutes and gently wash it by adding 600 &#181;L of MeOH and mixing gently to avoid breaking up the pellet</li>
		<li>Carefully remove the remaining MeOH and dry the protein pellet on a benchtop for approximately 5 min.</li>
		<li>(Optional) Perform an additional centrifugation step to spin down any broken pellet after the MeOH wash to avoid loss of sample. 
		NOTE: The experiment can be stopped after the CM precipitation step. Once the pellet is obtained, it may be stored in 500 &#181;L of MeOH in -20 &#176;C up to a week.
		<ol>
			<li>Dissolve the protein pellet in 200 &#181;L of 2SHB buffer by vortexing at 42 &#176;C/1500 rpm in a thermal shaker until the pellet is dissolved.</li>
		</ol>
		</li>
		<li>(Optional) Incubate for an additional 5&#8211;10 min in a sonicating water bath to dissolve the pellet. 
		NOTE: The length of sonication varies depending on solubility of the material. However, prolonged sonication can cause protein degradation.</li>
	</ol>
	</li>
	<li>Prepare 0.2% methyl methanethiosulfonate (MMTS) (v/v) in 2SHB by adding 2 &#181;L of MMTS to 998 &#181;L of 2SHB buffer. 
	NOTE: Use freshly prepared 0.2% MMTS for each experiment.</li>
	<li>Add 200 &#181;L of 0.2% MMTS in 2SHB to each tube to a final concentration of 0.1% MMTS. Incubate for 15 min at 42 &#730;C with shaking at 1500 rpm in a thermal shaker.</li>
</ol>
3. Acyl-RAC: Hydroxylamine (HAM) cleavage and capture of S-acylated proteins
<ol>
	<li>Repeat 3-4x CM precipitations as described above to remove MMTS. Removal of MMTS can be estimated by the lack of distinct odor of MMTS. After each precipitation, dissolve the pellet in 100 &#181;L of 2SHB buffer by vortexing at 42 &#730;C/1500 rpm in a thermal shaker until the pellet dissolves, and then dilute with 300 &#181;L of Buffer A.</li>
	<li>After final precipitation, dissolve samples in 200 &#181;L of 2SHB buffer as described above and dilute with 240 &#181;L of Buffer A.</li>
	<li>In case of comparing changes in S-acylation in response to several treatment conditions, measure protein concentration again and proceed with equal amount of protein for each condition.</li>
	<li>Retain 40 &#181;L from each sample as an input control.</li>
	<li>Split samples into two equal parts of 200 &#181;L and mark tubes as &#8220;+ HAM&#8221; and &#8220;- HAM&#8221;. Add 50 &#181;L of freshly prepared neutral 2 M HAM (pH 7.0-7.5) to a final concentration of 400 mM to one of the tubes (+ HAM) and 50 &#181;L of neutral 2 M NaCl to the second tube (- HAM), which will be used as a negative control. Proceed to addition of thiopropyl-Sepharose (TS) beads. 
	NOTE: The experiment can be stopped after any of the CM precipitation steps. Once the pellet is obtained, it may be stored in 500 &#181;L of MeOH in -20 &#176;C up to a week. Neutral pH of 2 M HAM ensures its selectivity for the acyl-cysteine thioester bond and should be carefully adjusted. Care should be taken when handling samples in 2SHB buffer to avoid loss of sample due to excessive foaming. All following steps are identical for - HAM and + HAM samples.</li>
	<li>Add 30 &#181;L of TS bead-slurry to each tube and rotate the tubes for 1-2 h at RT.</li>
	<li>Wash the TS beads 4x with 1% SDS in Buffer A to remove residual HAM.
	<ol>
		<li>Gently spin down all bead samples using a microfuge for 1 min and carefully aspirate the supernatant.</li>
		<li>Resuspend the beads in 500 &#181;L of 1% SDS in Buffer A.</li>
		<li>Repeat step 3.7.1&#8211;3.7.2 thrice. 
		NOTE: Activated thiopropyl-Sepharose (TS) is supplied freeze-dried in the presence of additives that must be washed away at neutral pH before coupling. Distilled water is recommended for swelling and washing. Weigh 0.1 g of beads and resuspend in 1 mL of distilled water and allow it to swell while rotating for 30 min-1 h at RT. Wash beads 1x with buffer A and prepare a slurry with buffer A, in a ratio of 50% settled medium to 50% buffer. Swollen TS may be stored at neutral pH in the presence of 20% ethanol at 4 &#176;C up to a week. Do not use sodium azide as a bacteriostatic agent since azide ions react with the 2-pyridyl disulfide groups. For higher efficiency, prepare fresh beads. While handling the beads, the tip of a P200 pipette tip can be cut slightly so as to prevent any damage. 
		NOTE: The experiment can be stopped at any stage after CM precipitation. The pellet may be store in 500 &#181;L of MeOH in -20 &#176;C up to a week.</li>
	</ol>
	</li>
</ol>
4. Elution and detection of S-acylated proteins
<ol>
	<li>After the last wash, gently spin down the beads as described above and aspirate as much supernatant as possible without disturbing the beads.</li>
	<li>Recover the proteins from beads with 4x SDS sample buffer with DTT.
	<ol>
		<li>Add 50 &#181;L of 4x SDS sample buffer to the beads and incubate at 80 &#730;C, 1500 rpm for 15 min in a thermal shaker. Let the tubes cool.</li>
		<li>Centrifuge the beads at 5,000 x g for 3 min to completely pellet the beads and transfer the eluted proteins to a fresh 1.5 mL tube using a gel loading tip.</li>
	</ol>
	</li>
	<li>Run SDS-PAGE and analyze S-acylation of the protein(s) of interest by western blotting.</li>
</ol>

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No conflicts of interest declared.

Here, we successfully utilized the acyl-RAC assay to detect S-acylation of selected proteins in both cultured human cells and primary cells derived from mouse tissue. This method is simple, sensitive, and can be easily performed with minimal equipment requirements using standard biochemistry techniques. This method has been shown to successfully identify novel S-acylated proteins such as the &#946;-subunit of the protein translocating system (Sec61b), ribosomal protein S11 (Rps11), and microsomal glutathione-S-transferase 3 (MGST3)22. The sensitivity and adaptability of acyl-RAC makes it suitable to study protein S-acylation in a variety of biological samples.
A significant fraction of S-acylated proteins can be associated with glycosphingolipid-enriched parts of the plasma membrane, also known as lipid rafts. Thus, some S-acylated proteins might not be fully extracted if mild detergents, such as Triton X-100, NP-40 or Brij58, are used for lysis30. Anionic detergents (such as SDS) or maltoside based non-ionic detergents (such as n-dodecyl &#946;-D-maltoside (DDM)) can be used to ensure dissociation of lipid rafts and full recovery of the proteins of interest30,31,32. In this protocol, we used DDM, a mild, lipid-like non-ionic detergent that has been demonstrated to be effective in membrane protein extraction and maintaining their stable native state over prolonged periods33.
The unregulated thioesterase activity during lysis of the sample can potentially result in reduced protein yield after the TS pull-down step and thus affect the detection of S-acylated proteins. A thioesterase inhibitor, such as ML211, a dual inhibitor for Acyl-Protein Thioesterase 1 (APT1, LYPLA1) and Acyl-Protein Thioesterase 2 (APT2, LYPLA2) can be added to prevent indiscriminate deacylation of proteins in the lysate34,35.
An incomplete blockade of cysteine residues can cause binding of non-acylated proteins to TS, thus resulting in a high background evident by high signal in the &#8211;HAM samples. The background binding can be substantially reduced by an additional incubation with a thiol-crosslinking agent-2,2&#8217;-dithiodipyridine as described in a modified ABE protocol by Zhou et al.36.
In contrast to metabolic labeling, acyl-RAC is not limited to live cells and can be performed to detect protein S-acylation in freshly isolated or frozen tissue samples. Since the acyl-RAC protocol avoids an immunoprecipitation step, it can be used to simultaneously detect lipidation of multiple proteins of interest in the same assay, thus potentially reducing the amount of biological material required in a single experiment. However, metabolic labeling-based assays are more suitable for experiments aimed to specifically detect de novo S-acylation or measure fatty acid turnover rates on a protein of interest. Another important limitation of acyl-RAC is that this technique is unable to distinguish between different fatty acids that can be covalently bound to the cysteine residues via the same thioester bond3,4,5,6,7.
Both acyl-RAC and ABE assays are based on selective cleavage of the thioester bond between the cysteine residue and a fatty acid to reveal a free thiol group. Since acyl-RAC uses direct pull-down of S-acylated proteins by a thiol-reactive resin, this method requires fewer steps in comparison to ABE. Although similar, proteome-wide studies of S-acylation show some variation in the proteins detected using these two methods, most likely due to the technical differences. For example, in rat brain homogenate, ABE-based analysis identified 241 S-acylated proteins whereas acyl-RAC-based analysis identified 14425. 61 proteins in the rat brain proteome were commonly detected by both methods underscoring the need for use of multiple techniques to ascertain the S-acylation status of a protein.
Another potential drawback of acyl-RAC and ABE methods is the inability to reliably evaluate stoichiometry of S-acylation and determine the number of lipidated cysteines. A modification of these techniques, termed acyl-PEG exchange or PEG-shift assay, was developed to address these limitations37,38. In this method, pull-down step following hydroxylamine cleavage is substituted by incubation with a PEG-maleimide mass-tag. Resulting PEGylation of the exposed free cysteine residues can be observed as mass shift by SDS-PAGE.
In conclusion, acyl-RAC is a fast, sensitive and reliable method that can provide valuable insights into dynamics of protein S-acylation under both physiological and pathophysiological conditions in a great variety of biological samples. Considering limitations of the method discussed above, a combination of acyl-RAC with other techniques is recommended to fully characterize S-acylation of a protein of interest.

S-acylation is a reversible post-translational modification involving addition of a fatty acyl chain to an internal cysteine residue on a target protein via a labile thioester bond1. It was first reported as a modification of proteins with palmitate, a saturated 16-carbon fatty acid2, and therefore this modification is often referred to as S-palmitoylation. In addition to palmitate, proteins can be reversibly modified by a variety of longer and shorter saturated (myristate and stearate), monounsaturated (oleate) and polyunsaturated (arachidonate and eicosapentanoate) fatty acids3,4,5,6,7. In eukaryotic cells, S-acylation is catalyzed by a family of enzymes known as DHHC protein acyltransferases and the reverse reaction of cysteine deacylation is catalyzed by protein thioesterases, most of which still remain enigmatic8.
The lability of the thioester bond makes this lipid modification reversible, allowing it to dynamically regulate protein clustering, plasma membrane localization, intracellular trafficking, protein-protein interactions and protein stability9,10. Consequently, S-acylation has been linked to several disorders including Huntington&#8217;s disease, Alzheimer&#8217;s disease and several types of cancer (prostate, gastric, bladder, lung, colorectal), which necessitates development of reliable methods to detect this post-translational protein modification11.
Metabolic labeling with radioactive ([3H], [14C] or&#160;[125I]) palmitate was one of the first approaches developed to assay protein S-acylation12,13,14. However, radiolabeling-based methods present health concerns, are not very sensitive, time consuming, and only detect lipidation of highly abundant proteins15. A faster and nonradioactive alternative to radiolabeling is metabolic labeling with bioorthogonal fatty acid probes, which is used routinely to assay dynamics of protein S-acylation16. In this method, a fatty acid with a chemical reporter (alkyne or azide group) is incorporated into the S-acylated protein by a protein acyltransferase. Azide-alkyne Huisgen cycloaddition reaction (click chemistry)&#160;can then be used to attach a functionalized group, such as a fluorophore or biotin, to the integrated fatty acid allowing for detection of the S-acylated protein17,18,19.
Acyl-biotin exchange (ABE) is one of the extensively used biochemical methods for capture and identification of S-acylated proteins that bypasses some of the shortcomings of metabolic labeling such as unsuitability for tissue samples15. This method can be applied for analysis of S-acylation in a diverse range of biological samples, including tissues and frozen cell samples20,21. This method is based on selective cleavage of the thioester bond between the acyl group and the cysteine residue by neutral hydroxylamine. The liberated thiol groups are then captured with a thiol-reactive biotin derivative. The generated biotinylated proteins are then affinity-purified using streptavidin agarose and analyzed by immunoblotting.
An alternative approach termed acyl-resin assisted capture (Acyl-RAC) was later introduced to replace the biotinylation step with direct conjugation of free cysteines by a thiol-reactive resin22,23. This method has fewer steps compared to ABE and similarly can be used to detect protein S-acylation in a wide range of samples1.
Acyl-RAC consists of 4 main steps (Figure 1), 
1. Blocking of free thiol groups; 
2. Selective cleavage of the cysteine-acyl thioester bond with neutral hydroxylamine (HAM) to expose cysteine thiol groups; 
3. Capturing of the lipidated cysteines with a thiol-reactive resin; 
4. Selective enrichment of the S-acylated proteins after elution with reducing buffer.
The captured proteins can then be analyzed by immunoblotting or subjected to mass spectrometry (MS) based proteomics to assess the S-acylated proteome in a varied range of species and tissues22,24,25. Individual S-acylation sites can also be identified by trypsin digestion of the captured proteins and analysis of the resulting peptides by LC-MS/MS22. Here, we demonstrate how acyl-RAC can be used for simultaneous detection of S-acylation of multiple proteins in both a cell line and a tissue sample.

<table><tbody><tr><td>cOmplete Protease Inhibitor Cocktail tablets</td><td>Sigma</td><td>11836170001</td><td></td></tr><tr><td>Eppendorf Centrifuge 5424</td><td>Eppendorf</td><td>22620444</td><td></td></tr><tr><td>Hydroxylamine (HAM)</td><td>Sigma</td><td>159417</td><td></td></tr><tr><td>Methyl methanethiosulfonate (MMTS)</td><td>Sigma</td><td>64306</td><td></td></tr><tr><td>Mini tube rotator</td><td>LabForce</td><td></td><td></td></tr><tr><td>ML211</td><td>Cayman</td><td>17630</td><td></td></tr><tr><td>Multi-Therm Cool-Heat-Shake</td><td>Benchmark Scientific</td><td>H5000-HC</td><td></td></tr><tr><td>n-Dodecyl &amp;beta;-D-maltoside (DDM)</td><td>Sigma</td><td>D641</td><td></td></tr><tr><td>Phosphatase Inhibitor Cocktail 2</td><td>Sigma</td><td>P5726</td><td></td></tr><tr><td>Thiopropyl-Sepharose 6B (TS)</td><td>Sigma</td><td>T8387</td><td></td></tr><tr><td>Ultrasonics Quantrex Sonicator</td><td>L &amp;amp; R</td><td></td><td></td></tr></tbody></table>

detection of protein s-acylation using acyl-resin assisted capture

Protein S-acylation, also referred to as S-palmitoylation, is a reversible post-translational modification of cysteine residues with long-chain fatty acids via a labile thioester bond. S-acylation, which is emerging as a widespread regulatory mechanism, can modulate almost all aspects of the biological activity of proteins, from complex formation to protein trafficking and protein stability. The recent progress in understanding of the biological function of protein S-acylation was achieved largely due to the development of novel biochemical tools allowing robust and sensitive detection of protein S-acylation in a variety of biological samples. Here, we describe acyl resin-assisted capture (Acyl-RAC), a recently developed method based on selective capture of endogenously S-acylated proteins by thiol-reactive Sepharose beads. Compared to existing approaches, Acyl-RAC requires fewer steps and can yield more reliable results when coupled with mass spectrometry for identification of novel S-acylation targets. A major limitation in this technique is the lack of ability to discriminate between fatty acid species attached to cysteines via the same thioester bond.

Following the protocol described above, we first used acyl-RAC to simultaneously detect S-acylation of several proteins in Jurkat cells, an immortalized T cell line originally derived from the peripheral blood of a T cell leukemia patient27. Regulatory T cell proteins previously identified as S-acylated9,28,29 were chosen to demonstrate the utility of this method. As shown in Figure 2A, tyrosine kinase Lck can be detected in lysates treated with neutral 2 M hydroxylamine to cleave the thioester bond between cysteine residues and a fatty acid moiety (+ HAM lane). The sample treated with 2 M NaCl (- HAM lane) represents an important negative control, indicating the selectivity of the assay for detection of the thioester bonds. The input lane further confirms the specificity of the signal and can serve as a positive control if the protein of interest is not S-acylated. The membrane was then stripped and reprobed with antibodies against proteins Fyn and LAT to demonstrate that the acyl-RAC assay can be used to analyze S-acylation of multiple proteins at the same time (Figure 2A).
To demonstrate the utility of this method to detect protein S-acylation in tissue samples, we applied the acyl-RAC protocol to the mouse spleen. As shown in Figure 2B, S-acylation of Lck, Fyn and LAT can be readily detected in primary mouse splenocytes indicating that this modification is conserved between two species.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61016/61016fig1.jpg" /> 
Figure 1. Schematic representation of acyl-RAC. The method consists of 4 main steps: (1) blocking free thiol groups with methyl methanethiosulfonate (MMTS), (2) selective cleavage of the cysteine-acyl thioester bond with neutral hydroxylamine (HAM) to expose cysteine thiol groups, (3) capture of proteins with newly exposed cysteine thiol by direct conjugation with a thiol-reactive resin and (4) recovery of captured proteins using a reducing elution buffer, followed by immunoblotting or MS analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61016/61016fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61016/61016fig2.jpg" /> 
Figure 2. Detection of S-acylated proteins. An acyl-RAC assay was used to detect S-acylation of T cell signaling proteins in Jurkat T cells (A) and murine spleen tissue (B). Immunoblotting with Lck-, Fyn- and LAT-specific antibodies shows S-acylation of these proteins in the hydroxylamine treated sample (+ HAM lane). A sample treated with sodium chloride (- HAM lane) serves as negative control. Untreated lysates are shown in the input lane. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61016/61016fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Buffer</td>
			<td>Composition</td>
			<td>Notes</td>
		</tr>
		<tr>
			<td>Lysis Buffer (LB)</td>
			<td>1% DDM in DPBS; 10 &#181;M ML211; Phosphatase Inhibitor Cocktail 2 (1:100); Protease Inhibitor Cocktail (1X), PMSF (10 mM)</td>
			<td>Prepare fresh, chill at 4 &#730;C before use.</td>
		</tr>
		<tr>
			<td>SDS Buffer (2SHB)</td>
			<td>2% SDS; 5 mM EDTA; 100 mM HEPES; pH 7.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Buffer A</td>
			<td>5 mM EDTA; 100 mM HEPES; pH 7.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydroxylamine (HAM)</td>
			<td>2 M stock solution in distilled H20</td>
			<td>Prepare fresh. When first dissolved, HAM has a very low pH. Use 5 M NaOH to pH to 7-7.5.</td>
		</tr>
		<tr>
			<td>4X SDS sample buffer</td>
			<td>200 mM Tris-Cl (pH 6.8), 8% SDS (sodium dodecyl sulfate) 0.01% Bromophenol blue, 10% glycerol</td>
			<td>Supplement with 5 mM DTT just before use.</td>
		</tr>
	</tbody>
</table>
Table 1: Buffer composition of commonly used buffers required in the protocol. 2SHB buffer and Buffer A can be prepared in advance but their pH must be checked regularly. Lysis buffer and HAM must be prepared the day of experiment.

Watch this Scientific Journal Video about Detection of Protein S-Acylation using Acyl-Resin Assisted Capture at JoVE.com

Detection of Protein S-Acylation using Acyl-Resin Assisted Capture

Acyl-RAC (Acyl-Resin Assisted Capture) is a highly sensitive, reliable and easy to perform method to detect reversible lipid modification of cysteine residues (S-acylation) in a variety of biological samples.

Protein S-acylation, also referred to as S-palmitoylation, is a reversible post-translational modification of cysteine residues with long-chain fatty ...

detection-of-protein-s-acylation-using-acyl-resin-assisted-capture

Nanyang Technological University

Research

JoVE Journal

Biochemistry

9.9K Views.  McGovern Medical School at UT Health. Acyl-hars assisted capture of Acyl-RAC is een gevoelige en betrouwbare methode die kan worden gebruikt om eiwit S-acyleratie te detecteren in een verscheidenheid van biologische monsters. Dit protocol omzeilt enkele van de beperkingen van metabole etikettering en radio-etikettering, en maakt de gelijktijdige detectie van S-acylation van meerdere eiwitten. Niet alleen levende cellen, maar ook primaire weefsels en bevroren monsters.S-acylation kan een verscheidenheid aan cellulaire proce...

Video: Detection of Protein S-Acylation using Acyl-Resin Assisted Capture

Deacetylation Assays to Unravel the Interplay between Sirtuins (SIRT2) and Specific Protein-substrates

Acetylation has emerged as an important post-translational modification (PTM) regulating a plethora of cellular processes and functions. This is further supported by recent findings in high-resolution mass spectrometry based proteomics showing that many new proteins and sites within these proteins can be acetylated. However the identity of the enzymes regulating these proteins and sites is often unknown. Among these enzymes, sirtuins, which belong to the class III histone lysine deacetylases, have attracted great interest as enzymes regulating the acetylome under different physiological or pathophysiological conditions. Here we describe methods to link SIRT2, the cytoplasmic sirtuin, with its substrates including both in vitro and in vivo deacetylation assays. These assays can be applied in studies focused on other members of the sirtuin family to unravel the specific role of sirtuins and are necessary in order to establish the regulatory interplay of specific deacetylases with their substrates as a first step to better understand the role of protein acetylation. Furthermore, such assays can be used to distinguish functional acetylation sites on a protein from what may be non-regulatory acetylated lysines, as well as to examine the interplay between a deacetylase and its substrate in a physiological context.

Deacetylation Assays to Unravel the Interplay between Sirtuins SIRT2 and Specific Protein-substrates

This protocol describes the required steps to execute in vitro and in vivo deacetylation assays in order to establish the role of proteins as specific deacetylation substrates for sirtuins and further study the role of reversible - lysine acetylation as a post-translational modification.

Acetylation has emerged as an important post-translational modification (PTM) regulating a plethora of cellular processes and functions. This is further ...

Chemistry

Utilizing a Comprehensive Immunoprecipitation Enrichment System to Identify an Endogenous Post-translational Modification Profile for Target Proteins

It is now well-appreciated that post-translational modifications (PTMs) play an integral role in regulating a protein&#39;s structure and function, which may be essential for a given protein&#39;s role both physiologically and pathologically. Enrichment of PTMs is often necessary when investigating the PTM status of a target protein, because PTMs are often transient and relatively low in abundance. Many pitfalls are encountered when enriching for a PTM of a target protein, such as buffer incompatibility, the target protein antibody is not IP-compatible, loss of PTM signal, and others. The degree of difficulty is magnified when investigating multiple PTMs like acetylation, ubiquitination, SUMOylation 2/3, and tyrosine phosphorylation for a given target protein. Studying a combination of these PTMs may be necessary, as crosstalk between PTMs is prevalent and critical for protein regulation. Often, these PTMs are studied in different lysis buffers and with unique inhibitor compositions. To simplify the process, a unique denaturing lysis system was developed that effectively isolates and preserves these four PTMs; thus, enabling investigation of potential crosstalk in a single lysis system. A unique filter system was engineered to remove contaminating genomic DNA from the lysate, which is a problematic by-product of denaturing buffers. Robust affinity matrices targeting each of the four PTMs were developed in concert with the buffer system to maximize the enrichment and detection of the endogenous states of these four PTMs. This comprehensive PTM detection toolset streamlines the process of obtaining critical information about whether a protein is modified by one or more of these PTMs.

Investigating multiple, endogenous post-translational modifications for a target protein can be extremely challenging. Techniques described here utilize an optimized lysis buffer and filter system developed with specific PTM-targeting, affinity matrices to detect the acetylation, ubiquitination, SUMOylation 2/3, and tyrosine phosphorylation modifications for a target protein using a single, streamlined system.

It is now well-appreciated that post-translational modifications (PTMs) play an integral role in regulating a protein&#39;s structure and function, which ...

Detection of Protein Ubiquitination Sites by Peptide Enrichment and Mass Spectrometry

The posttranslational modification of proteins by the small protein ubiquitin is involved in many cellular events. After tryptic digestion of ubiquitinated proteins, peptides with a diglycine remnant conjugated to the epsilon amino group of lysine (&#39;K-&#949;-diglycine&#39; or simply &#39;diGly&#39;) can be used to track back the original modification site. Efficient immunopurification of diGly peptides combined with sensitive detection by mass spectrometry has resulted in a huge increase in the number of ubiquitination sites identified up to date. We have made several improvements to this workflow, including offline high pH reverse-phase fractionation of peptides prior to the enrichment procedure, and the inclusion of more advanced peptide fragmentation settings in the ion routing multipole. Also, more efficient cleanup of the sample using a filter-based plug in order to retain the antibody beads results in a greater specificity for diGly peptides. These improvements result in the routine detection of more than 23,000 diGly peptides from human cervical cancer cells (HeLa) cell lysates upon proteasome inhibition in the cell. We show the efficacy of this strategy for in-depth analysis of the ubiquitinome profiles of several different cell types and of in vivo samples, such as brain tissue. This study presents an original addition to the toolbox for protein ubiquitination analysis to uncover the deep cellular ubiquitinome.

We present a method for the purification, detection, and identification of diGly peptides that originate from ubiquitinated proteins from complex biological samples. The presented method is reproducible, robust, and outperforms published methods with respect to the level of depth of the ubiquitinome analysis.

The posttranslational modification of proteins by the small protein ubiquitin is involved in many cellular events. After tryptic digestion of ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Neuroscience

Optimized Incorporation of Alkynyl Fatty Acid Analogs for the Detection of Fatty Acylated Proteins using Click Chemistry

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in addition to its implications in cancer and neurodegenerative diseases. Recent developments in fatty acylation detection methods have enabled efficient and non-hazardous detection of fatty acylated proteins, particularly through the use of click chemistry with bio-orthogonal labeling. However, click chemistry detection can be limited by the poor solubility and potential toxic effects of adding long chain fatty acids to cell culture. Described here is a labeling approach with optimized delivery using saponified fatty acids in combination with fatty-acid free BSA, as well as delipidated media, which can improve detection of hard to detect fatty acylated proteins. This effect was most pronounced with the alkynyl-stearate analog, 17-ODYA, which has been the most commonly used fatty acid analog in click chemistry detection of acylated proteins. This modification will improve cellular incorporation and increase sensitivity to acylated protein detection. In addition, this approach can be applied in a variety of cell types and combined with other assays such as pulse-chase analysis, stable isotope labeling with amino acids in cell culture, and mass spectrometry for quantitative profiling of fatty acylated proteins.

The study of fatty acylation has important implications in cellular protein interactions and diseases. Presented here is a modified protocol to improve click chemistry detection of fatty acylated proteins, which can be applied in various cell types and combined with other assays, including pulse-chase and mass spectrometry.

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in ...

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

Histone Proteomic Analysis and Post-Translational Modification Characterization

Histone Modification Screening using Liquid Chromatography, Trapped Ion Mobility Spectrometry, and Time-Of-Flight Mass Spectrometry

Histone proteins are highly abundant and conserved among eukaryotes and play a large role in gene regulation as a result of structures known as posttranslational modifications (PTMs). Identifying the position and nature of each PTM or pattern of PTMs in reference to external or genetic factors allows this information to be statistically correlated with biological responses such as DNA transcription, replication, or repair. In the present work, a high-throughput analytical protocol for the detection of histone PTMs from biological samples is described. The use of complementary liquid chromatography, trapped ion mobility spectrometry, and time-of-flight mass spectrometry (LC-TIMS-ToF MS/MS) enables the separation and PTM assignment of the most biologically relevant modifications in a single analysis. The described approach takes advantage of recent developments in dependent data acquisition (DDA) using parallel accumulation in the mobility trap, followed by sequential fragmentation and collision-induced dissociation. Histone PTMs are confidently assigned based on their retention time, mobility, and fragmentation pattern.

Author Spotlight: Enhanced Histone PTM Isomer Identification Through LC-TIMS-ToF MS/MS and PASEF

An analytical workflow based on liquid chromatography, trapped ion mobility spectrometry, and time-of-flight mass spectrometry (LC-TIMS-ToF MS/MS) for high confidence and highly reproducible "bottom-up" analysis of histone modifications and identification based on principal parameters (retention time [RT], collision cross section [CCS], and accurate mass-to-charge [m/z] ratio).

Histone proteins are highly abundant and conserved among eukaryotes and play a large role in gene regulation as a result of structures known as ...

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry (CCMS)

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity chromatography 1 or Activity Based Protein Profiling 2. Trifunctional Capture Compounds (CCs, Figure 1A) 3 are the basis for a generic approach, in which the initial equilibrium-driven interaction between a small molecule probe (the selectivity function, here S-adenosyl-L-homocysteine, SAH, Figure 1A) and target proteins is irreversibly fixed upon photo-crosslinking between an independent photo-activable reactivity function (here a phenylazide) of the CC and the surface of the target proteins. The sorting function (here biotin) serves to isolate the CC - protein conjugates from complex biological mixtures with the help of a solid phase (here streptavidin magnetic beads). Two configurations of the experiments are possible: "off-bead" 4 or the presently described "on-bead" configuration (Figure 1B). The selectivity function may be virtually any small molecule of interest (substrates, inhibitors, drug molecules).
S-Adenosyl-L-methionine (SAM, Figure 1A) is probably, second to ATP, the most widely used cofactor in nature 5, 6. It is used as the major methyl group donor in all living organisms with the chemical reaction being catalyzed by SAM-dependent methyltransferases (MTases), which methylate DNA 7, RNA 8, proteins 9, or small molecules 10. Given the crucial role of methylation reactions in diverse physiological scenarios (gene regulation, epigenetics, metabolism), the profiling of MTases can be expected to become of similar importance in functional proteomics as the profiling of kinases. Analytical tools for their profiling, however, have not been available. We recently introduced a CC with SAH as selectivity group to fill this technological gap (Figure 1A).
SAH, the product of SAM after methyl transfer, is a known general MTase product inhibitor 11. For this reason and because the natural cofactor SAM is used by further enzymes transferring other parts of the cofactor or initiating radical reactions as well as because of its chemical instability 12, SAH is an ideal selectivity function for a CC to target MTases. Here, we report the utility of the SAH-CC and CCMS by profiling MTases and other SAH-binding proteins from the strain DH5&alpha; of Escherichia coli (E. coli), one of the best-characterized prokaryotes, which has served as the preferred model organism in countless biochemical, biological, and biotechnological studies. Photo-activated crosslinking enhances yield and sensitivity of the experiment, and the specificity can be readily tested for in competition experiments using an excess of free SAH.

Profiling of Methyltransferases and Other S-adenosyl-L-homocysteine-binding Proteins by Capture Compound Mass Spectrometry CCMS

Capture Compounds are trifunctional small molecules to reduce the complexity of the proteome by functional reversible small molecule-protein interaction followed by photo-crosslinking and purification. Here we use a Capture Compound with S-adenosyl-L-homocysteine-binding as selectivity function to isolate methyltransferases from an Escherichia coli whole cell lysate and identify them by MS.

There is a variety of approaches to reduce the complexity of the proteome on the basis of functional small molecule-protein interactions such as affinity ...

Biology

A Facile Protocol to Generate Site-Specifically Acetylated Proteins in Escherichia Coli

Post-translational modifications that occur at specific positions of proteins have been shown to play important roles in a variety of cellular processes. Among them, reversible lysine acetylation is one of the most widely distributed in all domains of life. Although numerous mass spectrometry-based acetylome studies have been performed, further characterization of these putative acetylation targets has been limited. One possible reason is that it is difficult to generate purely acetylated proteins at desired positions by most classic biochemical approaches. To overcome this challenge, the genetic code expansion technique has been applied to use the pair of an engineered pyrrolysyl-tRNA synthetase variant, and its cognate tRNA from Methanosarcinaceae species, to direct the cotranslational incorporation of acetyllysine at the specific site in the protein of interest. After first application in the study of histone acetylation, this approach has facilitated acetylation studies on a variety of proteins. In this work, we demonstrated a facile protocol to produce site-specifically acetylated proteins by using the model bacterium Escherichia coli as the host. Malate dehydrogenase was used as a demonstration example in this work.

A Facile Protocol to Generate Site-Specifically Acetylated Proteins in Escherichia Coli

Genetic code expansion serves as a powerful tool to study a wide range of biological processes, including protein acetylation. Here we demonstrate a facile protocol to exploit this technique for generating homogeneously acetylated proteins at specific sites in Escherichia coli cells.

Post-translational modifications that occur at specific positions of proteins have been shown to play important roles in a variety of cellular processes. ...

Immunology and Infection

Quantification of Site-specific Protein Lysine Acetylation and Succinylation Stoichiometry Using Data-independent Acquisition Mass Spectrometry

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein functions, for example, by changing enzymatic activity or by mediating interactions. Precise quantification of site-specific protein acylation occupancy, or stoichiometry, is essential for understanding the functional consequences of both global low-level stoichiometry and individual high-level acylation stoichiometry of specific lysine residues. Other groups have reported measurement of lysine acetylation stoichiometry by comparing the ratio of peptide precursor isotopes from endogenous, natural abundance acylation and exogenous, heavy isotope-labeled acylation introduced after quantitative chemical acetylation of proteins using stable isotope-labeled acetic anhydride. This protocol describes an optimized approach featuring several improvements, including: (1) increased chemical acylation efficiency, (2) the ability to measure protein succinylation in addition to acetylation, and (3) improved quantitative accuracy due to reduced interferences using fragment ion quantification from data-independent acquisitions (DIA) instead of precursor ion signal from data-dependent acquisition (DDA). The use of extracted peak areas from fragment ions for quantification also uniquely enables differentiation of site-level acylation stoichiometry from proteolytic peptides containing more than one lysine residue, which is not possible using precursor ion signals for quantification. Data visualization in Skyline, an open source quantitative proteomics environment, allows for convenient data inspection and review. Together, this workflow offers unbiased, precise, and accurate quantification of site-specific lysine acetylation and succinylation occupancy of an entire proteome, which may reveal and prioritize biologically relevant acylation sites.

Here, we present unbiased quantification of site-specific protein acetylation and/or succinylation occupancy (stoichiometry) of an entire proteome through a ratiometric analysis of endogenous modifications to modifications introduced after quantitative chemical acylation using stable isotope-labeled anhydrides. In combination with sensitive data-independent acquisition mass spectrometry, accurate site occupancy measurements are obtained.

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein ...

Detection of Protein S-Acylation using Acyl-Resin Assisted Capture

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