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-t...

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.

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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 2...

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

Research

JoVE Journal

Biochemistry

9.8K Views.  McGovern Medical School at UT Health. 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.

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

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of programmed DNA breaks in lymphocytes. The Ataxia-Telangiectasia Mutated kinase (ATM), ATM- and Rad3-Related kinase (ATR) and the catalytic subunit of DNA-dependent Protein Kinase (DNA-PKcs) are among the first that are activated upon induction of DNA damage, and are central regulators of a network that controls DNA repair, apoptosis and cell survival. As part of a tumor-suppressive pathway, ATM and ATR activate p53 through phosphorylation, thereby regulating the transcriptional activity of p53. DNA damage also results in the formation of so-called ionizing radiation-induced foci (IRIF) that represent complexes of DNA damage sensor and repair proteins that accumulate at the sites of DNA damage, which are visualized by fluorescence microscopy. Co-localization of proteins in IRIFs, however, does not necessarily imply direct protein-protein interactions, as the resolution of fluorescence microscopy is limited. 
In situ Proximity Ligation Assay (PLA) is a novel technique that allows the direct visualization of protein-protein interactions in cells and tissues with unprecedented specificity and sensitivity. This technique is based on the spatial proximity of specific antibodies binding to the proteins of interest. When the interrogated proteins are within ~40 nm an amplification reaction is triggered by oligonucleotides that are conjugated to the antibodies, and the amplification product is visualized by fluorescent labeling, yielding a signal that corresponds to the subcellular location of the interacting proteins. Using the established functional interaction between ATM and p53 as an example, it is demonstrated here how PLA can be used in suspension cell cultures to study the direct interactions between proteins that are integral parts of the DNA damage response.

Here, it is demonstrated how the in situ Proximity Ligation Assay (PLA) can be used to detect and visualize the direct protein-protein interactions between ATM and p53 in suspension cell cultures exposed to genotoxic stress.

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of ...

mRNA Interactome Capture from Plant Protoplasts

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional gene regulation (PTGR) of messenger RNAs (mRNAs). In the past few years, a number of mRNA-bound proteomes from yeast and mammalian cell lines have been successfully isolated through the use of a novel method called &#34;mRNA interactome capture,&#34; which allows for the identification of mRNA-binding proteins (mRBPs) directly from a physiological environment. The method is composed of in vivo ultraviolet (UV) crosslinking, pull-down and purification of messenger ribonucleoprotein complexes (mRNPs) by oligo(dT) beads, and the subsequent identification of the crosslinked proteins by mass spectrometry (MS). Very recently, by applying the same method, several plant mRNA-bound proteomes have been reported simultaneously from different Arabidopsis tissue sources: etiolated seedlings, leaf tissue, leaf mesophyll protoplasts, and cultured root cells. Here, we present the optimized mRNA interactome capture method for Arabidopsis thaliana leaf mesophyll protoplasts, a cell type that serves as a versatile tool for experiments that include various cellular assays. The conditions for optimal protein yield include the amount of starting tissue and the duration of UV irradiation. In the mRNA-bound proteome obtained from a medium-scale experiment (107 cells), RBPs noted to have RNA-binding capacity were found to be overrepresented, and many novel RBPs were identified. The experiment can be scaled up (109 cells), and the optimized method can be applied to other plant cell types and species to broadly isolate, catalog, and compare mRNA-bound proteomes in plants.

Here, we present an interactome capture protocol applied to Arabidopsis thaliana leaf mesophyll protoplasts. This method critically relies on in vivo UV crosslinking and allows for the isolation and identification of plant mRNA-binding proteins from a physiological environment.

RNA-binding proteins (RBPs) determine the fates of RNAs. They participate in all RNA biogenesis pathways and especially contribute to post-transcriptional ...

Horizontal Gel Electrophoresis for Enhanced Detection of Protein-RNA Complexes

Native polyacrylamide gel electrophoresis is a fundamental tool of molecular biology that has been used extensively for the biochemical analysis of RNA-protein interactions. These interactions have been traditionally analyzed with polyacrylamide gels generated between two glass plates and samples electrophoresed vertically. However, polyacrylamide gels cast in trays and electrophoresed horizontally offers several advantages. For example, horizontal gels used to analyze complexes between fluorescent RNA substrates and specific proteins can be imaged multiple times as electrophoresis progresses. This provides the unique opportunity to monitor RNA-protein complexes at several points during the experiment. In addition, horizontal gel electrophoresis makes it possible to analyze many samples in parallel. This can greatly facilitate time course experiments as well as analyzing multiple reactions simultaneously to compare different components and conditions. Here we provide a detailed protocol for generating and using horizontal native gel electrophoresis for analyzing RNA-Protein interactions.

Native polyacrylamide gel electrophoresis is a fundamental tool for analyzing RNA-protein interactions. Traditionally most experiments have used vertical gels. However, horizontal gels provide several advantages, such as the opportunity to monitor complexes during electrophoresis. We provide a detailed protocol for generating and using horizontal native gel electrophoresis.

Native polyacrylamide gel electrophoresis is a fundamental tool of molecular biology that has been used extensively for the biochemical analysis of ...

Detection of Heterodimerization of Protein Isoforms Using an in Situ Proximity Ligation Assay

Regulated protein-protein interactions are a guiding principle for many signaling events, and the detection of such events is an important element in understanding how such pathways are organized and how they function. There are many methods to detect protein-protein interactions in cells, but relatively few can be used to detect interactions between endogenous proteins. One such method, the proximity ligation assay (PLA), has several advantages to recommend its use. Compared to other common methods of protein-protein interaction analysis, PLA has relatively high sensitivity and specificity, can be performed with minimal cell manipulation, and, in the protocol described herein, requires only two target-specific antibodies derived from different species (e.g., from mouse and rabbit) and one specialized reagent: a set of secondary antibodies that are covalently linked to specific oligonucleotides that, when brought in close proximity of one another, create an amplifiable platform for in situ PCR or rolling circle amplification. In this presentation, we show how to apply the PLA technique to visualize changes in MST1 and MST2 proximity in fixed cells. The technique described in this manuscript is particularly applicable for the analysis of cell signaling studies.

Detection of Heterodimerization of Protein Isoforms Using an in Situ Proximity Ligation Assay

Here, we show how to use a Proximity Ligation Assay (PLA) to visualize MST1/MST2 heterodimerization in fixed cells with high sensitivity.

Regulated protein-protein interactions are a guiding principle for many signaling events, and the detection of such events is an important element in ...

Extracellular Protein Microarray Technology for High Throughput Detection of Low Affinity Receptor-Ligand Interactions

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling cascades during homeostasis and disease, and as such represent prime therapeutic targets. Despite their relevance, these interaction networks remain significantly underrepresented in current databases; therefore, most extracellular proteins have no documented binding partner. This discrepancy is primarily due to the challenges associated with the study of the extracellular proteins, including expression of functional proteins, and the weak, low affinity, protein interactions often established between cell surface receptors. The purpose of this method is to describe the printing of a library of extracellular proteins in a microarray format for screening of protein-protein interactions. To enable detection of weak interactions, a method based on multimerization of the query protein under study is described. Coupled to this microbead-based multimerization approach for increased multivalency, the protein microarray allows robust detection of transient protein-protein interactions in high throughput. This method offers a rapid and low sample consuming-approach for identification of new interactions applicable to any extracellular protein. Protein microarray printing and screening protocol are described. This technology will be useful for investigators seeking a robust method for discovery of protein interactions in the extracellular space.

Here, we present a protocol to screen extracellular protein microarrays for identification of novel receptor-ligand interactions in high throughput. We also describe a method to enhance detection of transient protein-protein interactions by using protein-microbead complexes.

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling ...

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture (CARIC) Strategy

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used strategy for RBP capture exploits the polyadenylation [poly(A)] of target RNAs, which mostly occurs on eukaryotic mature mRNAs, leaving most binding proteins of non-poly(A) RNAs unidentified. Here we describe the detailed procedures of a recently reported method termed click chemistry-assisted RNA-interactome capture (CARIC), which enables the transcriptome-wide capture of both poly(A) and non-poly(A) RBPs by combining the metabolic labeling of RNAs, in vivo UV cross-linking, and bioorthogonal tagging.

Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture CARIC Strategy

A detailed protocol for applying the click chemistry-assisted RNA-interactome capture (CARIC) strategy to identify proteins binding to both coding and noncoding RNAs is presented.

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used ...

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and the degradation of target proteins in live cells. By using short (pulse) radiolabeling times (&lt;30 min) and tightly controlled chase times, it is possible to label only a small fraction of the total protein pool and follow its folding. When combined with nonreducing/reducing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoprecipitation with (conformation-specific) antibodies, folding processes can be examined in great detail. This system has been used to analyze the folding of proteins with a huge variation in properties such as soluble proteins, single and multi-pass transmembrane proteins, heavily N- and O-glycosylated proteins, and proteins with and without extensive disulfide bonding. Pulse-chase methods are the basis of kinetic studies into a range of additional features, including co- and posttranslational modifications, oligomerization, and polymerization, essentially allowing the analysis of a protein from birth to death. Pulse-chase studies on protein folding are complementary with other biochemical and biophysical methods for studying proteins in vitro by providing increased temporal resolution and physiological information. The methods as described within this paper are adapted easily to study the folding of almost any protein that can be expressed in mammalian or insect-cell systems.

Here we describe a protocol for a general pulse-chase method that allows the kinetic analysis of folding, transport, and degradation of proteins to be followed in live cells.

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and ...