Chemical-proteomic strategies to investigate cysteine posttranslational modifications

Biochimica et Biophysica Acta 1844 (2014) 2315–2330

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Review

Chemical-proteomic strategies to investigate cysteine posttranslational modifications Shalise M. Couvertier, Yani Zhou, Eranthie Weerapana ⁎ Boston College, Chestnut Hill, MA 02467, USA

a r t i c l e

i n f o

Article history: Received 30 May 2014 Received in revised form 8 September 2014 Accepted 29 September 2014 Available online 5 October 2014 Keywords: Oxidation Nitrosation Palmitoylation Prenylation Glutathionylation Lipid-derived electrophile

a b s t r a c t The unique combination of nucleophilicity and redox-sensitivity that is characteristic of cysteine residues results in a variety of posttranslational modifications (PTMs), including oxidation, nitrosation, glutathionylation, prenylation, palmitoylation and Michael adducts with lipid-derived electrophiles (LDEs). These PTMs regulate the activity of diverse protein families by modulating the reactivity of cysteine nucleophiles within active sites of enzymes, and governing protein localization between soluble and membrane-bound forms. Many of these modifications are highly labile, sensitive to small changes in the environment, and dynamic, rendering it difficult to detect these modified species within a complex proteome. Several chemical-proteomic platforms have evolved to study these modifications and enable a better understanding of the diversity of proteins that are regulated by cysteine PTMs. These platforms include: (1) chemical probes to selectively tag PTM-modified cysteines; (2) differential labeling platforms that selectively reveal and tag PTM-modified cysteines; (3) lipid, isoprene and LDE derivatives containing bioorthogonal handles; and (4) cysteine-reactivity profiling to identify PTM-induced decreases in cysteine nucleophilicity. Here, we will provide an overview of these existing chemical-proteomic strategies and their effectiveness at identifying PTM-modified cysteine residues within native biological systems. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Cysteine residues play critical roles within protein scaffolds as sites of nucleophilic and redox catalysis, allosteric regulation, metal binding and structural stabilization [1]. These diverse functions of cysteine are facilitated by the unique nucleophilic and redox properties of the cysteine thiol group. Due to the low dissociation energy of the S-H bond and the large atomic radius of sulfur, the cysteine thiol has a pKa value of ~ 8.0 [2]. This side-chain pKa is easily perturbed by the local protein environment, with pKa values as low as 3.5 reported for members of the glutaredoxin family [3,4]. These altered pKa values serve to enhance the nucleophilicity and redox sensitivity of thiols. This increased reactivity thereby promotes a multitude of electrophilic and oxidative posttranslational modifications (PTMs) (Fig. 1). Cysteine PTMs can be categorized into spontaneous and enzymecatalyzed modifications. Spontaneous cysteine PTMs are generally driven by the encounter of reactive cysteines with endogenous oxidants and reactive electrophiles, such as reactive oxygen/nitrogen species (ROS/RNS) and lipid-derived electrophiles (LDEs). Oxidative modifications of cysteine result in the formation of sulfenic/sulfinic/sulfonic acids [5], S-nitrosothiols [6], intra- and inter-chain disulfides [7], persulfides [8] and mixed disulfides with glutathione and cysteine [9]. LDEs such as ⁎ Corresponding author. E-mail address: [email protected] (E. Weerapana).

http://dx.doi.org/10.1016/j.bbapap.2014.09.024 1570-9639/© 2014 Elsevier B.V. All rights reserved.

4-hydroxy-2-nonenal (HNE) and 15-deoxy-Δ12,14-prostaglandin J2 (15d-PGJ2), commonly form Michael adducts with nucleophilic cysteines [10]. Enzyme-catalyzed modifications of cysteine include prenylation [11] and palmitoylation [12], whereby select enzymes mediate the transfer of the lipid or isoprenoid moiety from an activated donor to the thiol side chain of cysteine. It is important to note that some modifications, e.g. disulfide formation, can be both spontaneous and enzyme catalyzed. In addition to the common eukaryotic PTMs discussed above, many other rare modifications of cysteine, such as methylation and phosphorylation, have been discovered in both eukaryotic and prokaryotic organisms [13]. All of these cysteine PTMs have been shown to modulate protein activity as well as localization, and dysregulation of these PTMs are critical in driving a variety of proliferative and degenerative diseases [14–16]. Methods to enrich, identify and quantify cysteine PTMs are of paramount importance to our understanding of the scope and physiological role of these modifications. Chemical-proteomic approaches that investigate these modifications within the context of a native proteome have been developed over several decades [17]. These methods range from early radioisotope-labeling experiments to more recent advances in chemical-probe design and mass spectrometry (MS), which enable detailed interrogation of these modifications. The recent advent of bioorthogonal reactions that allow for selective tagging of functional groups with minimal perturbations to biomolecules [18] have further accelerated studies into cysteine PTMs. Here we will discuss the technologies currently available to investigate the common cysteine PTMs.

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OH S O Sulfinic acid

O R1 R Lipid-derived electrophile (LDE) adducts S

S

OH

Sulfenic acid

O Sulfonic acid

S SR

O S

OH S O

R= H glutathione cysteine protein thiol

SH

14

S-palmitoylation

S-sulfhydration S-glutathionylation S-cysteinylation inter/intramolecular disulfide

S NO

S n = 2 or 3 S-prenylation

n

S-nitrosation

Fig. 1. Common posttranslational modifications of cysteines.

Advantages and limitations of each method and biological insight that has been gained through utilization of these methods will be highlighted where relevant.

methods outlined in the cysteine oxidation section can also be used to detect these other oxidative cysteine PTMs. 2.1. Two-dimensional gel electrophoresis (2DE)

2. Cysteine oxidation Cysteine residues are highly susceptible to oxidation, resulting in the formation of a variety of cysteine oxoforms [19]. Of these, the most common is intra- and inter-molecular disulfide formation, which is known to facilitate protein folding and structural stability. In addition to disulfide bonds, other oxoforms of cysteine include sulfenic, sulfinic and sulfonic acids (Fig. 2A) [5]. Sulfenic acids are generated by the reaction of a cysteine thiol with biological oxidants such as hydrogen peroxide, hypochlorous acid and hydroxyl radicals, as well as the hydrolysis of S-nitrosothiols [19]. Further oxidation of sulfenic acids result in sulfinic and sulfonic acids. Sulfinic acids are stable to most cellular reductants, and are reduced only in the presence of a newly discovered ATPdependent sulfiredoxin enzyme [20]. Sulfonic acids are the most highly oxidized thiol species and are considered to be irreversible. The reversibility of cysteine sulfenic acids has resulted in the evolution of this PTM into a key mode of regulation in biology, with functions analogous to phosphorylation, acetylation and ubiquitination [21]. Proteins from diverse functional classes have been identified to be sulfenylated and sulfinylated, and regulate key signaling pathways in (patho)physiology [5]. Examples include the protein-tyrosine phosphatases (PTPs), whereby sulfenylation of the active-site cysteine nucleophile is a wellcharacterized mechanism for inactivation of these enzymes [22]. The formation of the sulfenic acid is subsequently followed by either disulfide formation with an adjacent cysteine, or sulfonamide formation with a backbone amide-nitrogen [23,24]. Similarly, cysteine proteases such as the deubiquitinating enzymes (DUBs) that cleave ubiquitin from target proteins, are known to be sulfenylated at the active-site cysteine nucleophile. In particular, sulfenylation of both USP1, a DUB involved in DNA damage response [25], and A20, a known tumor suppressor [26], are known to inhibit deubiquitinase activity. These and other studies into the function of cysteine oxoforms have been accelerated by the availability of methods to identify these modifications. Although S-glutathionylation and S-nitrosation are described in separate sections of this review, it is important to note that many of the

The oxidation of cysteine residues into the various cysteine oxoforms serves to quench the nucleophilicity of the cysteine thiol. Existing two-dimensional gel electrophoresis (2DE) methods rely on wellcharacterized cysteine-reactive agents such as iodoacetamide (IAM) or N-ethylmaleimide (NEM) to cap all reduced cysteines in the proteome, leaving behind oxidized cysteines that are unreactive to the capping agent. These oxidized cysteine residues are then reduced with well-known reducing agents such as dithiothreitol (DTT) or tris(2carboxyethyl)phosphine (TCEP), and capped with IAM and NEM derivatives containing a radiolabel [27] or fluorophore [28,29] for detection. The resulting proteome samples are subject to 2DE and imaged by autoradiography or fluorescence to visualize oxidized proteins [30]. DTT and TCEP treatment results in reduction of numerous cysteine oxoforms [31], including sulfenic and sulfinic acids, nitrosothiols, and disulfides, thereby allowing for the facile visualization of multiple oxoforms of cysteine within a biological sample. However, this method relies on the initial cysteine capping step proceeding to completion; reduced efficiency in this step often results in false positives. Furthermore, comprehensive visualization of all oxidized proteins are limited by the resolution of 2DE, which results in co-elution of proteins with similar physicochemical properties, as well as the limit of detection afforded by the imaging methods, which hinders detection of low abundance oxidized proteins. To enable direct comparison of the degree of oxidation in two samples, differential labelling with orthogonal fluorophores in a method known as redox differential in-gel electrophoresis (redox-DIGE) was developed [32,33]. This method allows for facile visualization of differences in oxidized protein content across samples, but is limited by the inability to inform on the fraction of protein oxidized, since information on total protein levels is lacking. Differential labeling of reduced and oxidized cysteines with orthogonal fluorescent dyes helps to circumvent this problem by providing the fraction of oxidized protein [34]. However, this method loses the advantage of multiplexing, since each sample needs to be analyzed independently. A variety of methods exist for

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A H 2O 2, HOCl or, OH S

SH

OH

OH S O

OH S O Sulfinic acid

Sulfenic acid

O Sulfonic acid

B 12 C

12 C -ICAT

TCEP

S

1. Digest 2. LC/MS

13 C -ICAT

S[O]

S[O]

C

S

13 C

H

H H O

O

O

S

O

O O

S

O

+ H 2O H

H N O

O

O HN H

O

S

N3

N3 O

NH H

HO

DYn-1

E S O H

H

HO

H

S O

H

DAz-2

O

O

HO

DAz-1

O

O

OH

O

H

13 C

O

DCP-Bio1 O O

O

HO

S

m/z

OH

NH

H

12 C

O dimedone-cysteine adduct

DCP-FL1 O

S

O

O

-H+

dimedone

D

12 C

Intensity

S

SH

H

H BCN

Fig. 2. Cysteine oxidation. (A) Cysteine can be oxidized to sulfenic, sulfinic and sulfonic acids by reaction with hydrogen peroxide, perchloric acid or hydroxyl radicals. (B) OxICAT is a modified version of the ICAT technology geared to study cysteine oxidation. (C) The nucleophilic addition of dimedone to sulfenic acids generates a covalent adduct. (D) Dimedone-based probes for chemical-proteomics contain a biotin group for enrichment (DCP-Bio1), a fluorophore for imaging (DCP-FL1) or a bioorthogonal azide/alkyne (DAz-1, DAz-2, DYn-1). (E) A strained-alkyne probe, BCN, reacts with sulfenic acids to generate a covalent adduct.

differential labeling of proteomes with fluorophores for redox-DIGE experiments [35]. Each method requires stringent control of protein concentrations, labeling specificity and efficiency. The application of 2DE methods to identify cysteine oxidation events within a cellular milieu is highlighted in a study that utilized redox-DIGE to identify mitochondrial proteins sensitive to endogenous reactive oxygen species (ROS) generation [33]. A Cy3-NEM derivative was used to tag oxidized cysteines in control mitochondria, whereas proteins from redoxchallenged mitochondria were tagged with Cy5-NEM. Fluorescence visualization identified several proteins with increased oxidation under the redox-challenge. These proteins were identified by MS and two candidate proteins, propionyl-CoA carboxylase (PCC) and the pyruvate dehydrogenase kinase (PDHK), were selected for further characterization. The enzymatic activity of both of these proteins was shown to be sensitive to oxidative stress, thereby identifying two

novel redox-sensitive proteins in the mitochondria. A similar differential fluorescence labeling strategy was recently used to identify cysteines oxidized during acute hypoxia and subsequent reoxygenation [36]. This study identified the exact cysteine that is modified, thereby improving on previous techniques that only identify the oxidized protein. 2.2. Isotope-coded affinity tags (ICAT) Due to the poor reproducibility of 2DE and the low throughput involved in interfacing 2DE with MS analysis, where each spot on the gel has to be individually processed, a method to directly interface differential cysteine labeling with MS was developed. This method expands upon a shotgun proteomic strategy termed isotope-coded affinity tags (ICAT), whereby biotinylated-IAM derivatives containing

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isotopically light and heavy linkers are utilized [37]. Cysteines labeled with the ICAT reagents are enriched on streptavidin beads and analyzed by quantitative MS, whereby the tagged cysteine-containing peptides can be identified and the relative abundance between the light- and heavy-labeled samples can be quantified. The first iteration of ICAT in redox proteomics involved the use of the light and heavy ICAT reagents to compare reduced thiol content across two samples [38,39]. Using ICAT to compare a control proteome to one that is treated with hydrogen peroxide enabled the identification of cysteines sensitive to oxidation. Later iterations utilized ICAT to differentially label oxidized versus reduced cysteine residues within a single sample [40]. This method, termed OxICAT (Fig. 2B) allows for the quantification of the oxidized:reduced cysteine ratio, which can then be compared across numerous biological samples. Furthermore, to control for changes in protein levels, ICAT methods have been coupled with stable isotope dimethyl labeling to quantify protein abundance prior to cysteine enrichment [41,42]. Lastly, other labeling strategies have also been applied to incorporate stable isotopes for quantitative mass spectrometry. These include a method termed GELSILOX where stable oxygen isotopes are incorporated during the trypsin digestion step [43]. These isotopelabeled MS methods afford several advantages over 2DE, namely increased accuracy and quantification, global identification of all oxidized cysteines in a single experiment, as well as the ability to detect lower abundance proteins that are difficult to visualize with 2DE methods. However, this method relies on the cysteines of interest existing on tryptic peptides that are amenable to MS identification, thereby excluding cysteines located within very short or long tryptic peptides. Furthermore, as with the differential-tagging methods used for 2DE, OxICAT is subject to the limitation of requiring complete tagging of reduced cysteines to eliminate false positives. These ICAT-based methods have been applied to a variety of biological systems to identify oxidation-sensitive cysteines. In one such example, OxICAT was applied to interrogate oxidation-sensitive proteins in yeast [44]. Organelle fractionation followed by OxICAT determined the oxidation state of ~ 400 cysteines in control and hydrogen peroxide treated yeast cultures. One of the oxidation-sensitive proteins identified was GAPDH, where 75% of the active-site Cys150 was oxidized when exposed to a sub-lethal concentration of H2O2 for 15 minutes. This inactivation is likely responsible for the down-regulation of glycolysis upon exposure to oxidative stress, and redirection of glucose into the pentose phosphate pathway. The NADPH generated by the pentose phosphate pathway is an essential defense mechanism used by yeast for survival under oxidative conditions. 2.3. Chemical probes for sulfenic acids The 2DE and ICAT methods inform on all cysteine oxoforms, and do not differentiate between sulfenic acids, sulfinic acids and disulfides. Since sulfenic acids are transient and highly reversible, this modification, in particular, is attributed to mediate cellular signaling pathways akin to phosphorylation and acetylation. Methods to selectively identify sulfenic acids within a complex mixture are needed to better understand the targets, abundance and biological functions of these modifications. One of the early methods for modification of sulfenic acids relied on the selective reduction of sulfenic acids over other cysteine isoforms by arsenite, in a method analogous to the differential-tagging methods described previously [45]. However, sulfenic-acid specific chemical probes have been the most widely utilized platform for detection of this modification. These chemical probes include 5,5-dimethyl-1,3cyclohexanedione, commonly known as dimedone (Fig. 2C). Dimedone reactivity with sulfenic acids is driven by nucleophilic addition of the enolate intermediate from the 1,3-dicyclohexanedione moiety onto sulfenic acid to form a stable covalent adduct. Conjugation of dimedone or other dicarbonyl compounds to fluorophores and biotin (e.g. DCP-FL1 and DCP-Bio1 (Fig. 2D)) allow for visualization and avidin-enrichment for MS analysis [46–49]. Detailed investigations into the reactivity of

sulfenic acids and dimedone with commonly utilized thiol-capping agents such as IAM and NEM, have led to the recommendation that the dimedone addition be performed prior to thiol capping [50]. Limitations of biotin and fluorophore-tagged dimedone analogs arise from the bulky reporter tags that are unable to target buried sulfenic acids within proteins and additionally demonstrate poor cellpermeability. The lability of sulfenic acids necessitates methods to trap these modifications in vivo with minimal disruptions to local redox environments. To generate dimedone analogs that are able to trap sulfenic acids in live cells, derivatives with less sterically intrusive bioorthogonal handles were developed (Fig. 2D). These include azide-tagged probes (DAz-1 and DAz-2) [51,52] and an alkyne-tagged probe (DYn-1) [53] that can be conjugated to reporter tags after the initial protein labeling step by using a modified Staudinger reaction or copper-catalyzed azide-alkyne cycloaddition (CuAAC) chemistry. Each of these probes has been shown to be cell permeable and able to modify sulfenic acids within live cells. To quantify levels of sulfenic acids, isotope-coded dimedone and iododimedone (ICDID) were generated to react with oxidized and reduced cysteines respectively [54]. With these probes, the extent of sulfenic acid formation at a particular cysteine residue can be accurately quantified. In a similar strategy, lanthanide-chelating dimedone (LnDOTA-dimedone) and iodoacetamide (Ln-MeCAT-IAM) derivatives were applied together with inductively coupled plasma-MS (ICP-MS) to quantify sulfenic acid formation [55]. The Ln-DOTA-dimedone specifically modifies sulfenic acids, whereas the Ln-MeCAT-IAM reagent modifies reduced thiols. The presence of the lanthanide ion on probelabeled proteins allows for accurate quantification using ICP-MS. Successful quantification of sulfenic acids using these two methods was demonstrated on purified peptides and proteins. Due to the lack of an enrichment handle, these methods in their current form can only be applied to simplified systems and do not allow for quantification of sulfenic acids from a complex mixture. One limitation of the dimedone chemotype for sulfenic-acid labeling is the low reaction rate that necessitates the use of high millimolar concentrations of the probe for protein labeling. Recently, strained cycloalkynes, such as 9-hydroxymethylbicyclo[6.1.0]nonyne (BCN) (Fig. 2E), were shown to selectively modify sulfenic acids with reaction rates that exceed those of dimedone by over two orders of magnitude [56]. These probes can be utilized in cell lysates and for live-cell labeling at micromolar concentrations and add to the arsenal of chemical probes available to study protein sulfenylation. Dimedone-based probes have found widespread utility in identifying protein sulfenic-acids in a variety of biological systems. These dimedone-driven findings include the use of a biotin-functionalized dimedone to characterize the site of oxidation of the protein kinase Akt2 [57]. Oxidation of Cys124 is shown to inhibit Akt2 activity, thereby presenting a mechanism in which Akt2 is modulated concurrently by phosphorylation and cysteine oxidation. Biotin-functionalized dimedone was also applied to investigate cysteine sulfenylation implicated in T-cell activation [58]. These studies demonstrate that sulfenic-acid formation is an essential step in immune-cell signaling and function. Lastly, the alkyne-functionalized dimedone, DYn-2, enabled monitoring of global changes in protein sulfenylation upon epidermal growth factor receptor (EGFR)-mediated signaling [53]. Sulfenylation of EGFR itself, at Cys797, was shown to enhance kinase activity. Each of these examples serves to highlight the diverse functions of sulfenic acids in essential cellular signaling pathways. 3. Cysteine glutathionylation, cysteinylation and sulfhydration Glutathione (L-γ-glutamyl-L-cysteinylglycine) is primarily found in the cytosol in both a reduced (GSH) and oxidized form (GSSG), and can form mixed disulfides with cysteine residues on proteins in a process known as S-glutathionylation (Fig. 3A) [59,60].The formation of S-glutathionylated proteins is dependent on the GSH/GSSG ratio

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A

O

GSSG S SG + GSH

SH

O N H

HO NH 2

B O

SH H N

O

HO HN

N H

O

O

O OH

O

HN

O

H HN

N H

O

SH H N

O OH

O

O O

O O

H HN

NH S

SH H N

GSH

O

HO

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NH S

H

BioGSH

H

BioGEE

C SH

S NEM

S NEM

S SG

S SG

S H

NEM S

S

S

S

Grx GSH

S

S NEM-Biotin 1. NEM-Biotin 2. Enrichment

S

Fig. 3. Cysteine glutathionylation. (A) Glutathionylation is the formation of a mixed disulfide with glutathione (GSH). (B) Biotinylated glutathione adducts used to study glutathionylation include BioGSH and BioGEE. (C) Glutathionylated proteins can be identified by capping reduced cysteines with NEM followed by glutaredoxin (GRX)-mediated reduction of glutathionylated cysteines. The resulting newly-formed thiols are then capped with NEM-biotin for enrichment of glutathionylated proteins.

and involves a thiol/disulfide exchange between a protein thiol and GSSG and can also result from the reaction of S-nitrosated cysteines or sulfenic acids with glutathione [61].This process is highly reversible and removal of glutathionylated adducts is driven by thiol/disulfide exchange with free GSH, or enzymatically by glutaredoxin. Several important cellular pathways are regulated through glutathionylation. These include the NF-κB survival pathway, whereby S-glutathionylation of a critical DNA-binding cysteine of NF-κB inhibits DNA binding and downstream transcriptional activation [62]. Other key proteins in this pathway such as IκB kinase (IKK) are also known to be glutathionylated [63], rendering the NF-κB response system highly sensitive to oxidative stress and changes in cellular GSH/GSSG ratios. Diverse proteins such as protein kinases, phosphatases and ion channels have been shown to be glutathionylated [60], thereby warranting chemical-proteomic methods to identify and quantify this modification within complex proteomes [64]. S-cysteinylation is analogous to S-glutathionylation; although less characterized in eukaryotic systems it has been shown to be an important mediator of protein function in prokaryotes [29]. Lastly, cysteine sulfhydration results from the reaction of cysteine thiols with hydrogen sulfide (H2S), which is a critical mediator of cell signaling [65]. The resulting persulfide (-S-SH) shows reactivity similar to thiol groups resulting in challenges in developing detection methods [66–68]. Due to a scarcity of existing methods to specifically identify sites of Scysteinylation and sulfhydration, we will focus here on existing methods to identify S-glutathionylated cysteines, since these have been more widely utilized within a biological context. 3.1. Radiolabeled glutathione Initial methods for studying glutathionylation utilized radiolabeled GSH either in vitro by treatment of lysates with 3H or 35S-labeled GSH,

or in situ by metabolic labeling with 35S-cysteine [64,69]. For in situ analysis, incorporation of the radiolabel into glutathione is facilitated by the endogenous glutathione-synthesis machinery (γ-glutamylcysteine synthetase and glutathione synthetase). These cellular studies require treatment with cycloheximide to inhibit cellular protein synthesis, thereby ensuring that the radiolabeled cysteine can only be incorporated into proteins through S-glutathionylation. After exposing cells to oxidative stress, S-glutathionylated proteins can be separated by SDS-PAGE and imaged by autoradiography. The 35S-GSH radiolabeling method has been widely utilized and enabled many of the early discoveries including identification of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as a target of S-glutathionylation [70]. Major limitations of this method are the requirements for specialized training for use of radioactivity, the need to inhibit global protein synthesis, which disrupts regular cellular function, the potential of false positives resulting from other forms of S-thiolation such as S-cysteinylation, and the lack of an enrichment handle to isolate modified proteins. 3.2. Biotinylated glutathione To overcome several of the limitations associated with the use of radiolabeled glutathione, biotinylated glutathione analogs have been developed. Biotinylated glutathione is easily synthesized in vitro using the water-soluble biotinylation reagent, sulfosuccinimidyl-6(biotinamido)-hexanoate (sulfo-NHS-biotin), which couples biotin to the free 1° amine group of GSH or GSSG in amine-free buffer to form BioGSH (Fig. 3B) or BioGSSG [71]. In addition to GSH and GSSG, glutathione ethyl ester can also be biotinylated in a similar fashion to afford a cell-permeable GSH source, biotin-labeled glutathione ethyl ester (BioGEE) (Fig. 3B) [72]. After enrichment of biotinylated proteins on streptavidin beads, the bound proteins can be eluted by treatment

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with reducing agents to cleave the mixed disulfide bond between the protein and the biotinylated-GSH, followed by subsequent gel or MS analysis. These biotinylated glutathione derivatives have been utilized both in vitro and in cells to identify S-glutathionylated proteins. One such example is the use of BioGEE to identify proteins that are glutathionylated following treatment of cells with carbon monoxide (CO) [73]. CO is known to generate low levels of ROS through inhibition of cytochrome c oxidase, thereby indirectly perturbing the GSH/GSSG ratio and increasing protein glutathionylation. Using BioGEE, p65 has been identified as a target of glutathionylation and this PTM blocks NF-κB-p65 nuclear translocation. Although biotinylated glutathione analogs serve to overcome several of the limitations associated with the radiolabeled glutathione, a significant drawback is the presence of the bulky biotin group that may artificially perturb the subset of proteins targeted by this modification. Additionally, similar to the radiolabeled glutathione method, this method does not allow for identification of native glutathionylation events since it requires the addition of exogenous glutathione to the sample being analyzed. 3.3. Bioorthogonal glutathione analogs To overcome the limitations associated with the addition of exogenous glutathione to the sample under analysis, a recent method synthesized a GSH variant containing an azide bioorthogonal handle directly in cells [74]. This was achieved by mutation of glutathione synthetase (GS), which catalyzes coupling of γGlu-Cys to azido-Ala to form azidoGSH. Transfection of this mutant GS enabled the in situ generation of azido-GSH for protein labeling and detection using click chemistry. This elegant method was utilized in cells to monitor the increase in S-glutathionylation in the presence of hydrogen peroxide [74]. 3.4. Anti-glutathione antibodies Antibodies for glutathione have been generated for the identification of glutathionylated proteins by immunoblotting or enrichment by immunoprecipitation [75]. These antibodies enable the interrogation of endogenous glutathionylated proteins. However, problems arise from the poor specificity of these antibodies, resulting in crossreaction with other epitopes within a proteome. Due to the small size of the glutathione epitope and the flexibility, detection is affected by the local environment at the site of glutathionylation. Another limiting factor is the low sensitivity of detection, which biases the method toward identification of the most abundant targets of glutathionylation [64]. A similar method exploits the affinity of a glutathione Stransferase (GST) from Schistososma japonicum to specifically bind to glutathionylated proteins [76]. In this GST-overlay method, protein mixtures are separated by SDS-PAGE, transferred onto nitrocellulose membranes and exposed to biotin-labeled GST. Glutathionylated proteins can be visualized by chemiluminescence after exposure to horseradish peroxidase (HRP)-avidin. This method has not been widely utilized since the initial report, and therefore the specificity and sensitivity are poorly characterized. 3.5. GRX-mediated reduction Glutaredoxins (GRX) are thioredoxin-family members that are known to reduce glutathione-mixed disulfides [77]. In the GRX-mediated reduction method, proteome samples are treated with cysteine-reactive agents such as NEM to cap reduced cysteines, followed by treatment with GRX to selectively reduce glutathionylated cysteines [78]. The newly revealed thiols are then labeled with a biotinylated thiolreactive agent, enabling selective enrichment of glutathionylated proteins and peptides (Fig. 3C). This method was used to identify glutathionylated proteins in ECV304 cells upon treatment with diamide to induce oxidative stress [78]. This method suffers from the same

limitations associated with the differential-labeling methods described previously, in that false positives can arise from incomplete cysteine capping in the first step of the process. 4. Cysteine nitrosation The conversion of cysteine thiols to S-nitrosothiol (SNO) is termed cysteine nitrosation [79,80]. Although the term S-nitrosylation is also commonly used to refer to this modification, S-nitrosation is the proper term for formation of a nitroso group (R-NO) from NO [79]. SNO synthesis in cells can result from three distinct pathways (Fig. 4A): (1) reaction of cysteine thiols with N2O3 resulting from autooxidation of nitric oxide (NO); (2) recombination of NO with thiyl radicals; and, (3) transition metal-catalyzed addition of NO to a thiol. The extent of SNO formation is governed by cellular nitric oxide (NO) levels, which are in turn regulated by three isoforms of NO synthase (NOS), which oxidize L-arginine to afford NO and L-citrulline [81]. Protein SNO formation can occur by transnitrosation via transfer of a NO group from low-molecular weight SNOs (e.g. S-nitrosocysteine (CysNO)/S-nitrosoglutathione (GSNO)) or other proteins [82]. To date, over 3000 sites of S-nitrosation have been identified on cellular proteins [83], yet no sequence motif or pKa trend has been observed across these identified sites of nitrosation [84,85]. These identified sites of nitrosation include active-site cysteines in proteins like caspase-3 (CASP3) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH). In CASP3, nitrosation of the active-site cysteine has been shown to inhibit protease activity, and the balance between cellular nitrosating and denitrosating activity governs the initiation of apoptosis [86,87]. GAPDH is similarly nitrosated on the active site Cys150, and nitrosation results in increased binding to the ubiquitin ligase Siah1, which facilitates GAPDH translocation into the nucleus [88,89]. The identification and characterization of these and other targets of S-nitrosation were enabled by chemical-proteomic methods to enrich and identify SNO-modified proteins from complex proteomes [90]. The low abundance and poor stability of endogenous SNO adducts have rendered the direct enrichment and identification of these modified species a challenge. Early methods for studying S-nitrosation included UV-visible and chemiluminescence detection, as well as photolysis to homolytically cleave S-N bonds, followed by detection of liberated NO [91,92]. These methods can quantify total nitrosothiol content in a sample, but do not allow for direct visualization or identification of individual S-nitrosated proteins. Here, we highlight several commonly utilized proteomic strategies that can enrich, identify and differentiate between S-nitrosated protein species [93]. 4.1. Biotin-switch technique The biotin-switch technique (BST) is the most widely utilized method for protein-SNO detection. The original BST includes three basic steps (Fig. 4B): (1) cap free cysteine thiols with the cysteine alkylating agent methyl methanethiosulfonate (MMTS); (2) selectively reduce SNOs to free thiols using ascorbate; and, (3) label the newly-formed thiols with the biotinylating cysteine-reactive agent N-[6-(biotinamido) hexyl]-3’-(2’-pyridyldithio)propionamide (biotin-HPDP) [94,95]. Biotinylated proteins can be detected by immunoblotting, or subjected to avidin enrichment for analysis by MS. Since the original report of BST, several iterations of this method have been reported [96,97], including the enrichment of biotinylated peptides instead of proteins (SNOSID) [98] and the use of resin-assisted capture (SNO-RAC) [99]. BST has also been interfaced with existing quantitative proteomic methods, such as ICAT, stable isotope labeling by amino acids in cell culture (SILAC), and tandem mass tags (TMT), to render the ability to identify and quantify sites of endogenous protein nitrosation [100–104]. These methods involve the incorporation of an isotopic label through metabolic labeling with heavy amino acids prior to BST, or labeling of reactive amino-acid side chains with isotopically labeled tags after the BST protocol. In addition to biotinylation of S-nitrosated proteins, a similar

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A

NO 2 N 2O3

O2 Reactive nitrogen SH species

S NO

RS R

RS

NO

RSH transition metal

R R

S

N O

S

N O

S

N O

e-

B SH

S

S

S

S

S

Biotin S S

S

N

ascorbate S NO (2)

S H

S

S

S

S

S

S

S

S

S NO

MMTS (1)

S

S

(3)

Biotin

O

C

N H PPh 2

S

R

R

S

S

PPh 3

R'

O

disulfide formation

sulfenamide (2)

(1)

X

R'

PPh 2

(3)

(2) X=O (3) X=S

O R

S

(1) N PPh 3

R

S

N O

R

N R' H "traceless" ligation

(4)

thiol-aza-ylide (6)

R'

S

(5) S S

O (4) X=O

X

(5) X=S PPh 2

R O

S P R SO3Na TXPTS adduct

3

N R' P Ph bis-ligation

(6)

P

Ph

SO3Na

3

Fig. 4. Cysteine nitrosation. (A) Nitrosation is the transfer of a nitroso-group onto cysteine residues, and can occur through three key pathways: reaction with N2O3, reaction with thiyl radicals, or transition-metal catalyzed addition of NO to thiols. (B) The biotin-switch technique (BST) involves capping of reduced thiols with MMTS, followed by selective reduction of nitrosated cysteines with ascorbate. The resulting newly-formed thiols are then capped with biotin-HPDP for enrichment of nitrosated proteins. (C) Chemical probes for nitrosation rely on the use of phosphine esters to form: (1) thio-aza-ylides; (2) sulfenamides; (3) traceless disulfides; (4) traceless sulfonamides; (5) bis-ligation products and (6) adducts with water-soluble TXPTS.

differential labeling approach has been taken to fluorescently label S-nitrosated cysteines to be compatible with 2DE analysis [105,106]. Despite the wide application of BST, several limitations should be noted. Primarily, as with the differential labeling methods used for cysteine oxidation, BST necessitates complete capping of free thiols in the initial alkylation step; incomplete alkylation will result in false positives. Furthermore, the lengthy sample preparation process increases the likelihood of SNO decomposition and scrambling of sites of S-nitrosation through disulfide exchange. The selectivity of ascorbate for reducing nitrosated cysteines relies on a mechanism postulated as an indirect reduction, whereby ascorbate does not directly donate

electrons to the nitrosothiol, but instead undergoes a transnitrosation reaction [107]. This selectivity for SNO has been brought into question by observations of disulfide reduction under these conditions [108]. Furthermore, several ascorbate-dependent artifacts have been reported to lead to false positives [109]. In response to the observed nonspecificity reported for BST, Stamler et al. showed that the ascorbatemediated reduction was highly specific, but inadvertent exposure to sunlight was the cause of false positives [110]. It was suggested that analysis of a sample in which all SNO species were photolyzed prior to processing can act as an additional control to reduce false positives in BST experiments [110].

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The advent of BST has enabled numerous biological studies that have illustrated the ubiquitous role of cysteine nitrosation in cellular function. A recent study utilized BST in red blood cells (RBCs) to show that NO generated by NOS is critical to maintain deformability in RBCs [111]. The cellular targets of NO in RBCs included several cytoskeleton proteins, such as α- and β-spectrins. Nitrosation of these cytoskeletal proteins were shown to be important to maintaining RBC deformation. Similarly, BST was critical for identifying proteins that mediate the NO-induced resistance of melanoma cells to anticancer agents such as cisplatin [112]. Several nitrosated proteins were identified, including caspase-3 and prolyl-hydroxylase-2, which are both inhibited by S-nitrosation. S-nitrosation of these and other proteins is thought to be directly involved in generating resistance to cisplatin. 4.2. Organomercury enrichment A method similar to BST involves the use of an organomercury resin (MRC) to enrich S-nitrosothiols [85]. This method exploits the reaction of phenylmercury with SNO to form stable thiol-mercury conjugates. The MRC resin is generated by coupling p-amino-phenyl mercuric acetate to N-hydroxysuccinimide (NHS)-functionalized agarose. Free cysteines within the sample are capped with MMTS, and S-nitrosated proteins are selectively captured on the MRC resin and subsequently released using performic acid, which concurrently oxidizes newly revealed cysteine thiols to sulfonic acids. The eluted peptides are then analyzed by MS, where the mass shift of the sulfonic acid modification enables identification of the exact site of S-nitrosation. This method was initially used to identify 328 nitrosated cysteines in mouse liver [85]. Further systematic analysis of various mouse tissues using MRC uncovered 1011 SNO sites in 647 proteins across mouse liver, brain, heart, kidney and lung [113]. These identified S-nitrosated proteins participate in diverse metabolic processes including glycolysis, gluconeogenesis, oxidative phosphorylation, amino-acid metabolism and the tricarboxylic acid cycle. In particular, S-nitrosation of Cys238 in very long chain acyl-coenzyme A dehydrogenase (VLCAD) was shown to activate the enzyme, thereby illustrating the role of S-nitrosation in regulating fatty-acid beta oxidation. Recently, BST was combined with MRC in a protein microarray platform. BST analysis was performed on a GSNO-pretreated high-density protein microarray chip containing 16,368 unique GST-tagged fulllength human proteins [114]. S-nitrosated sites were biotinylated using BST and detected using an anti-biotin antibody coupled with a fluorescent secondary antibody. Of the 834 S-nitrosated proteins identified, a subset was expressed and purified and subject to MRC enrichment and subsequent MS analysis to identify the sites of Snitrosation. The combining of orthogonal methods like BST and MRC to identify S-nitrosation provides increased confidence in the identified targets and serves to mitigate the limitations associated with each of the methods. 4.3. Chemical probes for nitrosothiols Due to the limitations and artifacts associated with BST analysis, significant efforts have been invested in developing chemical probes with selectivity for nitrosothiols over other cysteine oxoforms (Fig. 4C). Analogous to the Staudinger ligation [115], where phosphines react selectively with azides to generate aza-ylides, nitrosothiols can react with triarylphosphines to form thiol-aza-ylides [116]. The use of phosphine esters in place of phosphines resulted in rearrangement of the aza-ylide intermediate to form a sulfenamide [116]. Further engineering resulted in a ‘traceless’ SNO ligation with the use of phosphine ester or thioester derivatives [117], and the formation of a stable bisligation disulfide product [118,119]. Without bulky phosphine adducts, the disulfide products are more suitable for downstream analytical platforms. To circumvent the poor water solubility of these phosphine reagents, water-soluble tris(4,6-dimethyl-3-sulfonatophenyl)phosphine

trisodium salt hydrate (TXPTS) was utilized to modify nitrosothiols and form the corresponding aza-ylide. TXPTS reactivity with an S-nitrosated mutant of the alkylhydroperoxide reductase C (AhpC-SNO) was shown to proceed with high efficiency and the resulting covalent Salkylphosphonium product was stable and detectable by MS. [120] To further improve on the aqueous compatibility and robustness of these reactions, Tannenbaum and coworkers developed a panel of derivatized triphenylphosphines with sulfonate esters and tertiary amines to facilitate water solubility and MS ionization [121]. These modified probes were utilized to measure cellular GSNO levels using multiple reaction monitoring (MRM)-MS in malignant cancer cells and macrophages. These studies were the first to demonstrate the successful application of phosphine probes for nitrosothiols within a complex biological sample, however, only low molecular-weight thiols were analyzed. There are no reports on the application of phosphine probes to enrich and identify protein nitrosothiols from a complex proteome. With continued effort, the rates of reaction, water solubility of the phosphine reagent, stability of the resulting adducts and chemo-selectivity of the phosphine-nitrosothiol reaction will likely improve and enable the application of these probes to complex biological samples. 5. Cysteine prenylation Protein S-prenylation is the addition of an isoprenoid, either farnesyl (C15) or geranylgeranyl (C20), from the corresponding isoprenylpyrophosphate to a cysteine residue to form a thioether linkage [11]. These modifications are enzymatically installed by farnesyl transferase (FTase) or geranylgeranyltransferase (GGTase-I/II) (Fig. 5A). Both classes of transferases recognize the characteristic amino-acid motif at the C-terminus of prenylated proteins, known as a CaaX box. The CaaX motif contains the prenylated cysteine followed by two small aliphatic amino acids and an amino acid X that determines the type of isoprene conjugated to the protein. Some of the most studied targets of prenylation are the Ras and Rab subfamily of small GTP-binding proteins, which are critical to cell signaling during cell growth and differentiation [122,123]. Ras proteins are farnesylated at a CaaX motif and this modification is essential for oncogenic forms of Ras to transform cells. Instead of a CaaX box, Rab proteins contain C-terminal CC or CxC motifs where both cysteines are geranylgeranylated by GGTase-II. Prenylation of both Ras and Rab serves to localize these soluble proteins at the cell membrane. The requirement for CaaX, CxC or CC motifs render putative targets of prenylation amenable to identification through sequencemining methods. However, experimental methods that can identify and quantify prenylated proteins under different biological states will further our understanding of the ubiquity and functions of cellular prenylation [124–127]. 5.1. Radiolabeled isoprenes Early studies to identify endogenously prenylated substrates utilized radiolabeled isoprene derivatives as substrates for FTase and GGTases [128]. Treatment of cells with tritium 3H-labeled mevalonic acid resulted in the in situ generation of 3H-farnesyl and geranylgeranyl pyrophosphates. These radiolabeled isoprenes are then incorporated onto substrate proteins by FTase and GGTases and the resulting prenylated proteins can be visualized by SDS-PAGE and autoradiography. This method was critical to initial discoveries in the prenylation field, and demonstrated the substrate scope and abundance of this modification. For example, this method was applied in Schizosaccharomyces pombe, a variant of fission yeast, where the incorporation of 3Hmevalonate into 3H-farnesyl and geranylgeranyl pyrophosphates was demonstrated by HPLC. Autoradiography of the S. pombe proteome showed the prenylation of low molecular-weight proteins, likely to be GTP-binding proteins in this organism. These findings suggest that the fission yeast S. pombe, can be used as a model organism to study the underlying mechanistic basis of prenylation. In contrast, the budding yeast

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A

Prenyltransferase n = 2; FTase n = 3; GGTase

Prenyltransferase SH

S n = 2 or 3

Isoprenyl-pyrophosphate

2323

n

O

B HN O O P P O O O O O

O O P P O O O O O

H N n n = 1 or 2

(NH 4) 3

NO 2

HN O NH

NH H

H S

3

2

O

(NH 4) 3 BGPP

NBD-phosphoisoprenoid

C Lyse

Incubation with bio-orthogonal isoprenoid

HO

Streptavidin enrichment

SDS-PAGE

1.Digest 2.LC/MS

N3 Farnesyl azide N3 C15-dh-azide O

HO

HO

Incorporation of a fluorophore

alk-FOH

alk-FOH-2

Intensity

HO

Incorporation of biotin

m/z Fig. 5. Cysteine prenylation. (A) Cysteine prenylation is the conjugation of farnesyl or geranylgeranyl moieties to cysteines catalyzed by FTase or GGTase, respectively. (B) Fluorescent (NDB-phosphoisoprenoid) and biotin (BGPP) analogs of isoprenoid pyrophosphates have been developed for modification of prenylated proteins. (C) Bioorthogonal isoprenoids incorporate alkyne and azide groups to farnesol or geranylgeraniol. These probes are useful for identification of prenylated proteins in cells.

Saccharomyces cerevisiae, is unable to incorporate 3H-mevalonate due to poor uptake of this metabolic intermediate [129]. Radiolabeled methods to study prenylation are minimally invasive and do not require the chemical synthesis of modified isoprenes. However, limitations include the need to handle radiolabeled material, and the inability to use this method to enrich and identify unknown prenylated proteins from a complex mixture. 5.2. Fluorescent and biotinylated isoprenes To circumvent the use of radiolabels, fluorescent and biotin-tagged isoprene derivatives have been utilized for in-gel fluorescence and immunoblotting analyses of prenyltransferase activity. For fluorescent detection, 7-nitro-benzo[1,2,5]oxadiazol-4-ylamino (NBD) was conjugated to farnesyl and geranylgeranyl pyrophosphate (Fig. 5B), and shown to have affinities comparable to native substrates for FTase

and GGTase-I [130]. These fluorescent isoprenyl pyrophosphates were utilized in in vitro assays with several GTPase substrates and afforded fluorescent labeling, indicating successful transfer of the NBD-isoprene to protein targets. This method was adapted to a bead-based analysis to allow for the high-throughput measurement of FTase and GGTase-I activity on specific substrates, thereby enabling the screening of inhibitors for these enzymes. These NBD-tagged isoprenes were also shown to be cell permeable, and able to fluorescently tag overexpressed K-Ras in living cells. A similar approach was developed using biotingeranylpyrophosphate (BGPP) (Fig. 5B) [131]. This substrate was utilized by RabGGTase, but was not efficiently transferred by FTase and GGTase-I. To enable transfer of BGPP by FTase and GGTase-I, protein engineering was utilized to generate mutants of these enzymes with the ability to transfer BGPP to known substrates. To identify putative substrates of RabGGTase, and the engineered FTase and GGTase, these enzymes were added to cell lysates together with the BGPP substrate.

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Biotinylated proteins were enriched and analyzed by MS to identify targets of prenylation and screen for inhibitors of prenylation. These methods circumvent the need to use radioactivity, and provide powerful tools to study prenylation. However, limitations arise from the bulky fluorophore or biotin reporter groups, which show reduced affinity for the prenyltransferases and poor cell permeability.

5.3. Bioorthogonal isoprene reporters To overcome the limitations presented by the bulky reporter groups, less sterically invasive bioorthogonal reporter tags have been incorporated into isoprene substrates. Initial work in this area utilized azidefunctionalized farnesyl diphosphate and geranylgeranyl diphosphate analogs (Fig. 5C). Proteins modified by the azido-isoprenes were detected by incorporating a rhodamine or biotin tag using the Staudinger ligation or CuAAC [132,133]. Due to the high background labeling associated with CuAAC using azido-isoprenes and rhodamine-alkyne [134], alkyne-functionalized isoprene derivatives were also developed (Fig. 5C) [135]. These isoprene derivatives were shown to be cell permeable and allowed for visualization and identification of endogenously prenylated proteins. Since then, alkynyl-farnesol (alk-FOH) has been widely utilized as an efficient chemical reporter of S-prenylated proteins [136]. Use of the NBD and biotin-tagged analogs in proteomes required prior inhibition of endogenous isoprene production, whereas the alk-FOH method functions without depletion of cellular isoprenes. Use of alk-FOH has resulted in the discovery of previously uncharacterized sites of S-farnesylation, such as that of the zinc-finger antiviral protein (ZAP) [137]. Prenylation of ZAP is critical for targeting to endolysosomes and enhances antiviral activity. Furthermore, alk-FOH enabled the discovery that infection by the human pathogen Legionella pneumophila involves the prenylation of numerous Legionella proteins [138]. These prenylation events are important for localization of proteins to host organelles and are mediated by the host prenylation machinery. These studies highlight the application of bioorthogonal prenylation proteomics to provide insight into host-pathogen interactions. The advent of bioorthogonal isoprene derivatives has provided a powerful method to monitor S-prenylation both in vitro and in living cells. One possible limitation to this method is the possibility of off-target protein identifications due to metabolism of the isoprene derivatives into other lipid analogs when utilized in cells or in vivo.

A

6. Cysteine palmitoylation S-palmitoylation is the incorporation of palmitic acid onto cysteine residues of proteins (Fig. 6A) [139]. This posttranslational modification is enzymatically mediated by protein S-acyltransferases (PATs) containing a cysteine-rich domain with a conserved Asp-His-His-Cys (DHHC) signature motif. Numerous proteins have been identified to be palmitoylated, such as G proteins, ion channels and cytoskeletal proteins. Many of these are palmitoylated adjacent to a site of prenylation or myristoylation. Unlike S-prenylation, palmitoylation does not identify with a consensus motif, making this modification event difficult to predict by sequence analysis. Also unique to palmitoylation is the reversibility of the modification; this dynamic nature further complicates identification and analysis of endogenous palmitoylation events. Removal of palmitoyl groups is thought to be mediated by protein palmitoylthioesterase 1 (PPT1) in the lysosome, and acylprotein thioesterase 1 (APT1) in the cytoplasm. Similar to S-prenylation, early methods to study S-palmitoylation relied on metabolic labeling with radiolabeled lipid species. Exposing cells to 3H-palmitate resulted in incorporation of the radiolabeled lipid at sites of palmitoylation, enabling visualization with autoradiography [140]. These early methods are limited by the need to use large quantities of radioactive lipids and suffer from poor sensitivity, thereby spurring the development of alternative methods to study protein S-palmitoylation in cells [124–127]. Several targeted chemical-biology approaches have been utilized to study individual palmitoylated proteins, such as those from the Ras family [141,142], but here we will focus on proteomic methods that allow for the global and unbiased identification of protein palmitoylation within a complex biological system. 6.1. Acyl-biotin exchange To overcome the limitations of using radiolabeled palmitic acid to monitor S-palmitoylation, the acyl-biotin exchange (ABE) method was developed. This method relies on the sensitivity of the fatty acylthioester linkage to cleavage by hydroxylamine to generate the free cysteine thiol. The steps of the ABE method are (Fig. 6B): (1) capping of all free cysteine thiols with NEM; (2) selective cleavage of palmitoyl groups using hydroxylamine to reveal free thiols; and (3) conjugation of biotin to these newly exposed thiols using biotin-BMCC or biotin-HPDH

O

SH

O

C.

CoAS

N3

O

9

Acyltransferase

S

-azido-fatty acids n = 3, 5, 6, and 7

9

OH

n

O 17-ODYA

7

OH

B S

SH

NEM

NEM 1. Biotin-HPDP

O

O S

S NH 2OH

NEM

14

S

2. Enrichment 14

S

S-HPDP-Biotin

H

Fig. 6. Cysteine palmitoylation. (A) Cysteine palmitoylation is the enzymatic transfer of a palmitoyl group from palmitoyl-CoA to a cysteine residue. (B) The acyl-biotin exchange method uses NEM to cap free thiols, followed by removal of the palmitoyl group by NH2OH treatment. The resulting newly-formed thiols are then capped with biotin-HPDP for enrichment of palmitoylated proteins. (C) Bioorthogonal palmitate analogs incorporate either azide or alkyne groups on the carbon chain and can be used for identification of palmitoylated proteins in cells.

S.M. Couvertier et al. / Biochimica et Biophysica Acta 1844 (2014) 2315–2330

[143,144]. Biotinylated proteins can be subject to visualization by immunochemistry, or enrichment and identification with MS. A modified version of the ABE method is the S-acylation by resin-assisted capture (acyl-RAC) [145]. Free thiols generated after hydroxylamine treatment are captured on a thiol-reactive sepharose resin. ABE has facilitated several investigations into the biological role of S-palmitoylation. For example, MS analysis of human B lymphoid cells identified 95 putative palmitoylated proteins, including the two immune regulators, CD20 and CD23 [146]. The ABE method was interfaced with quantitative MS in a method termed palmitoyl-cysteine isolation capture and analysis (PICA) [147]. In PICA, palmitoylated species in two proteomes can be quantitatively compared by treating the free thiols generated from hydroxylamine treatment with either heavy or light ICAT reagents. Enrichment and MSanalysis of labeled peptides allows for identification and quantification of relative levels of palmitoylation from the two samples. This method was utilized to identify substrates for the different PAT enzymes, resulting in the characterization of CKAP4/p63 as a specific substrate for the PAT DHHC2 [147]. The ABE method has proved effective at enriching native palmitoylated proteins, but suffers from several limitations. Similar to the differential thiol labeling and biotin-switch techniques discussed previously, ABE relies on complete capping of free thiols with NEM for accuracy. Similarly, the poor specificity of hydroxylamine results in high background signals as well as identification of non palmitoylated proteins with thioester linkages, such as ubiquitin ligases and lipoamide-linked dehydrogenases. 6.2. Bioorthogonal palmitoylation reporters Metabolic labeling by a non-radioactive analog of palmitic acid allows for selective tagging of palmitoylated proteins in a method more direct than ABE. Several azide and alkyne-functionalized fatty acids (Fig. 6C) have been utilized as tools to tag endogenously palmitoylated proteins. Studies in this area were initiated by the synthesis of a series of ω-azido fatty acids of varying chain length [148]. These fatty-acid derivatives were evaluated in mammalian cells using the Staudinger ligation to conjugate a biotin group for visualization. Of the panel of ω-azido fatty acids, a 14-carbon chain analog was found to be selective for S-palmitoylation over N-myristoylation, another common lipid modification. Subsequent work identified the alkyne-functionalized lipid derivative, 17-octadecynoic acid (ODYA), as an alternative bioorthogonal label for S-palmitoylation [149]. ODYA is commercially available, cell permeable and readily incorporated into proteins in living cells. CuAAC was used to conjugate fluorescent and biotin tags for visualization and enrichment respectively. MS analysis of 17-ODYA treated Jurkat T cells resulted in the identification of ~125 putative palmitoylated proteins. In a similar study, a panel of ω-alkynyl fatty acids were synthesized and evaluated in mammalian cell culture for efficiency and selectivity of metabolic labeling of S-palmitoylated proteins [150]. Direct comparison of the azide and alkyne tags showed that the alkyne-functionalized lipid derivatives were more efficient for metabolic labeling compared to the azido analogs [151]. The advent of bioorthogonal lipid reporters for S-palmitoylation has enabled the discovery and biological function of novel protein targets of this modification. Metabolic labeling with 17-ODYA was interfaced with SILAC to provide a quantitative platform to monitor S-palmitoylation [152]. This SILAC platform was utilized to identify sites of palmitoylation that were enriched upon treatment with an inhibitor of palmitoyl protein thioesterases and to monitor turnover of palmitoylated proteins in cells using a pulse-chase labeling method with 17-ODYA and palmitic acid. Use of an alkynylated palmitic acid analog also facilitated the global profiling of S-palmitoylated proteins in a dendritic cell line [153]. These profiling studies identified IFITM3 as a palmitoylated protein, where palmitoylation is essential for membrane clustering of IFITM3 and antiviral activity. Other findings generated through these bioorthogonal labeling methods include: (1) the identification of flotillin-2 as a substrate of the PAT DHHC5; [154] (2) the demonstration that G

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protein-coupled receptor signaling is dependent on palmitoylation; [155] and (3) the implication of palmitoylation in regulation of meiotic entry in fission yeast [156]. Bioorthogonal reporters of palmitoylation offer high sensitivity, rapid detection and ability to enrich palmitoylated proteins from complex proteomes. However, these methods rely on the addition of a non-natural lipid moiety to cells at relatively high concentrations, which could potentially perturb the natural abundance and distribution of endogenous lipidated proteins. Further improvement is also needed in enabling visualization of lipidated proteins in living cells; the generation of live-cell compatible bioorthogonal chemistries could likely help with advancing this area of lipidated protein biology.

7. Cysteine adducts with lipid-derived electrophiles Lipid-derived electrophiles (LDEs) are products of cellular metabolism, cellular lipid nitrosation and peroxidation [10,157], the formation of which can be either non-enzymatic or enzymatic. Reactive oxygen species (ROS) and reactive nitrogen species (RNS) can trigger the non-enzymatic peroxidation of unsaturated fatty acids to afford α,βunsaturated aldehydes and nitroalkenes [158,159]. Examples include 4-hydroxy-2-nonenal (HNE) [160] and 2-trans-hexadecenal (2-HD) (Fig. 7A). Enzymatic LDE formation occurs in response to stimuli such as inflammation and include several prostaglandins such as 15-deoxyΔ12,13-prostaglandin J2 (15d-PGJ2), which are generated by the action of cyclooxygenases (COX), lipooxygenases (LOX), cytochrome P450s and NAD+/NADP+ -dependent dehydrogenases [161,162]. These LDEs share the common feature of being electron poor and able to react with nucleophilic cysteines via a Michael addition reaction (Fig. 7A). The resulting adducts are reversible and can regenerate the unmodified cysteine depending on the cellular conditions. Although LDEs can form adducts with histidine and lysine residues, preference has been observed for the reaction with the sulfhydryl group of cysteine [163]. The pKa of the cysteine thiol, its accessibility and the local steric and biological environment all serve to regulate the selectivity of cysteine LDE adducts [164,165]. LDE-modifications of proteins are known to regulate critical cellular functions. For example, Kelch-like ECH-Associate Protein 1 (Keap1) is a cysteine rich protein containing five functional domains. It acts as a cytoplasmic inhibitor of Nuclear Factor (Erythroid-derived-2)-like-2 (Nrf2) to prevent its translocation to the nucleus, where it binds to the antioxidant response elements (ARE) to activate ARE-dependent genes. Exposure of Keap1 to 15d-PGJ2 results in dissociation and nuclear translocation of Nrf2. Site-directed mutagenesis identified two Keap1 cysteines (Cys273 and Cys288) that were critical to this negative regulation. Thus, by modification of Keap1, LDEs can induce AREdependent genes for cytoprotection [166]. Heat shock proteins (HSPs) are another well-characterized target of LDEs. HSPs are molecular chaperones involved in protein trafficking and degradation, and are key mediators of the cellular stress response. In rats subject to a high fat/high-ethanol diet, both Hsp72 and Hsp90 have been shown to form HNE adducts at Cys267 and Cys572 respectively [167,168]. In both cases, modification by HNE inhibits the chaperone activity of the HSPs and results in impaired cellular stress response. To further our understanding of the endogenous targets of LDEmodification, numerous proteomic methods have been developed to visualize, enrich and identify LDE-modified proteins. Characterization of endogenous sites of LDE modification is limited by the low abundance of LDE-adducts, the reversibility of the Michael addition and potential side reactions generated by the carbonyl group present in the LDE adducts. Initial studies relied on the generation of monoclonal and polyclonal antibodies against LDE-amino acid adducts. These antibodies could be utilized for visualization and enrichment of LDE-modified proteins [165,169,170]. More recently, methods to selectively tag and enrich LDE-modified proteins have been developed and utilized.

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LDEs:

A

OH R

O

O

O

LDE

SH

* 2-HD

* HNE

S R

*

O

11

COOH *

O

15d-PGJ2

O

B

O H2N

O

O H2N

N H

N H

HN H

H N

NH H

C

OH O 3

Al-HNE

S O

O ARP

HN H

H N

OH O

NH H

3

N3

Az-HNE

S

O biotin hydrazide

DMSO

IAA

Heavy TEV tag N3

Intensity

D

m/z Identification heavy LDE

IAA

Light TEV tag N3

light Quantification

Fig. 7. Cysteine adducts with lipid-derived electrophiles (LDEs). (A) LDEs contain an alpha-beta unsaturated carbonyl that can react with cysteine residues via a Michael addition. Examples of common LDEs include HNE, 2-HD and 15d-PGJ2; * denotes carbon that is the site of nucleophilic attack. (B) Chemical derivatization of the carbonyl group that results from LDE-protein modification can be achieved using hydrazine (e.g. DNPH), amino-oxy (e.g. ARP) and hydrazide (e.g. biotin hydrazide) probes. (C) Azide and alkyne-tagged LDEs allow for labeling of LDEmodified proteins in living cells. (D) A competitive labeling platform allows for quantification of the extent of LDE labeling. Lysates are treated with an LDE and then subject to labeling with an iodoacetamide-alkyne (IAA) probe that labels reactive cysteines. LDE labeling is identified as a loss in cysteine reactivity, which is quantified through incorporation of isotopic labels for MS analysis.

7.1. Chemical derivatization The carbonyl group that is formed upon LDE modification of cysteines serves as a selective chemical-derivatization handle to enrich LDE targets [171]. This carbonyl group can be subject to trapping with hydrazines and hydrazides to form hydrazones that can be further reduced to a more stable adduct. Initial chemical-derivatization platforms for LDE-modified proteins used dinitrophenylhydrazine (DNPH) in a method known as an “Oxyblot”. Lysates treated with DNPH can be analyzed using Western blotting with an anti-DNPH antibody [172, 173]. A similar reagent, termed Girard’s P reagent, has also been utilized to tag carbonylated proteins [174]. Lastly, fluorescently labeled hydrazides have been used for facile visualization of protein carbonylation [175]. To incorporate an enrichment handle into these reagents to facilitate proteomic analyses, biotinylated aldehyde-capture reagents were developed. These include N’-aminooxymethylcarbonylhydrazino D-biotin (aldehyde-reactive probe, ARP) [176], and biotin hydrazide (Fig. 7B) [177]. Biotinylation of carbonylated proteins in cell lysates is achieved by treatment with either the hydroxylamine-based ARP or hydrazide probes. Treatment with the hydrazide probe requires sodium

borohydride-mediated reduction of the hydrazine bond prior to further processing; a step that is not necessary for the hydroxylamine-based ARP. The biotinylated proteins are captured on streptavidin resin and subject to MS analysis. ARP was used to identify HNE-modified proteins in rat heart mitochondria, resulting in the identification of the site of HNE-adduction in long-chain-specific acyl-CoA dehydrogenase [176]. Similarly, the hydrazide probe was used to identify HNE-adducts from RKO human colorectal carcinoma cells treated with HNE. This study provided a global survey of protein targets of HNE, and revealed the sensitivity of cellular pathways such as the proteasome and chaperonin systems to HNE-modification [177]. Further modifications to this platform include use of a hydrazide-functionalized, isotope-coded affinity tag (HICAT) to covalently label and quantify HNE-modified peptides [178]. HICAT comprises three moieties: a hydrazide functional group for covalent modification of carbonyl groups, an isotopically light or heavy succinic anhydride-derived linker for relative quantification, and a biotin group for enrichment of labeled peptides using an avidin affinity column. Application of HICAT to mitochondrial proteomes identified and quantified numerous HNE-modified proteins both in in vitro HNE-treated samples, as well as in cardiac mitochondria without

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addition of exogenous HNE [178]. Lastly, a comparative analysis of four hydrazine-based aldehyde-reactive probes (DNPH, biotin hydrazide (BH), ARP and a long-chain biotin hydrazide (LCBH)) showed that ARP and DNPH performed better than BH and LCBH, at least on a limited set of model peptides [170]. This study also confirmed that carbonyl modification of HNE-adducted peptides serves to prevent neutral loss of the HNE group during collision-induced dissociation (CID) fragmentation, thereby facilitating identification of the exact site of HNE modification. Further derivations of the biotin-hydrazide tagging approach include the use of a solid-phase hydrazide (SPH) reagent for enrichment and subsequent release of LDE-modified peptides using acid treatment [179]. This approach facilitates identification of the exact site of labeling by eliminating the biotin group prior to MS analysis. Quantification of the SPH enriched peptides has been achieved through 18O incorporation [180], stable isotope dimethyl labeling of the peptide amine groups [181], and d0/d4-succinic anhydride labeling [182]. These carbonyl derivatization methods have greatly advanced our knowledge of protein carbonylation during the last decade. However, limitations to these approaches lie in the promiscuity of hydrazine reagents that are known to react with all aldehydes, ketones and sulfenic acids on biomolecules [183], resulting in potential false-positive identifications of protein-LDE adducts. 7.2. Bioorthogonal reporters of LDE-adducts Biotinylated analogs of LDEs such as 15d-PGJ2 have been used to identify LDE-adducts in biological systems [184,185]. However, due to potential disruption of LDE-binding due to the presence of the biotin group, less intrusive functionalities are likely to be more advantageous of the detection of LDE adducts. Similar to the alkyne and azide-tagged isoprene and palmitoyl derivatives discussed previously, methods for identifying LDE-adducts have similarly exploited bioorthogonal tagging methods. Towards this end, azido-tagged HNE (Az-HNE) and alkynyltagged HNE (Al-HNE) (Fig. 7C) were synthesized and utilized to label proteins in intact RKO cells [186]. Az-HNE and Al-HNE adducted proteins were conjugated to a biotin reporter element using either the Staudinger ligation or CuAAC. MS analysis of RKO cells treated with the azide/alkyne-tagged HNE derivatives identified HNE-adduction of numerous stress-related proteins like HSPs at Az-HNE and Al-HNE concentrations as low as 5 μM [186]. The Al-HNE reporter was also used to identify HNE-modified peptides from healthy human plasma [187]. After treatment with Al-HNE, CuAAC was used to incorporate a photocleavable biotin linker for enrichment and subsequent release of HNE-adducted peptides for MS analysis. The use of the photocleavable linker minimized elution of non-specifically bound peptides from the streptavidin beads. Using this strategy, 18 sites of HNE modification were identified from human plasma including human serum albumin and apolipoprotein A1. The use of HNE-analogs bearing bioorthogonal reporter elements helps to overcome the non-specificity of the carbonyl-derivatization methods described previously. However, this method requires addition of an exogenous HNE derivative to cells or lysates and therefore does not allow for identification and quantification of endogenous HNE-modification sites. 7.3. Competitive labeling strategies To enable the quantification of the extent of HNE adduction of proteins within a complex proteome, a competitive chemicalproteomic strategy was developed (Fig. 7D) [188]. This method relies on the use of an iodoacetamide alkyne (IAA) probe to label reactive cysteines, coupled with a quantitative MS platform termed isoTOP-ABPP to quantify the extent of cysteine labeling [189]. Briefly, a cell lysate is treated with an LDE, followed by the IAA probe to cap all non-LDE modified reactive cysteines. The IAA-tagged proteins are conjugated to isotopically tagged cleavable linkers (TEV-light and TEV-heavy) for enrichment,

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selective release and MS-based identification and quantification of IAA-modified peptides. Comparison of an LDE-treated (TEV-light) sample to an untreated (TEV-heavy) sample allows for quantification of the extent of LDE-modification of each reactive cysteine in the proteome. In this study, MDA-MB-231 human breast cancer cells were treated with three LDEs (HNE, 15d-PGJ2 or 2-HD) and 750-1000 cysteines were quantified in terms of their sensitivities to each LDE. Several LDE-sensitive cysteines were identified, including a conserved, active site-proximal cysteine on ZAK kinase. HNE modification of this cysteine inhibits enzyme function and suppresses activation of the JNK signaling pathway during oxidative stress. The competitive isoTOP-ABPP platform is an indirect measure of cysteine modification by LDEs and is therefore susceptible to false positives generated by other cysteine modifications that perturb the nucleophilicity. Furthermore, observed changes could also be a reflection of variations in protein abundance across two independent biological samples. However, one important benefit of this platform is the ability to quantify the sensitivity of a cysteine towards a particular LDE, which was not possible with the previous methods. This method can also be extended to many of the other cysteine PTMs described in this review. 8. Summary Cysteine PTMs have been shown to be ubiquitous and highly dynamic, and therefore responsible for temporal and spatial regulation of diverse proteins. The dynamic nature of many of these modifications renders them a challenge to study within native biological systems. During the past decade, significant strides have been made toward developing analytical platforms to globally identify these cysteine PTMs and understand the functional role of these modifications. These methods are diverse and exploit the unique chemical reactivity of these modified species as handles to selectively enrich PTM-modified cysteines. In parallel, advances in MS instrumentation and quantification accuracy have further advanced this field by enabling the identification of sites of cysteine modification and quantification of the abundance of these PTMs. As these analytical platforms improve further, and our understanding of the reactivity of cysteine PTMs expands, the methods highlighted in this review are likely to improve in both selectivity and sensitivity. Until then, the many diverse techniques available for each type of cysteine PTM allow the use of multiple orthogonal methods to demonstrate and confirm the presence of a particular modification on a protein of interest. Given the diverse functional roles of reactive cysteines in human physiology, these proteomic methods will be critical to identifying dysregulated protein activities associated with disease. Acknowledgements Eranthie Weerapana is a Damon Runyon-Rachleff Innovator (DRR18-12). We are also grateful for financial support from the Smith Family Foundation and Boston College. We thank members of the Weerapana Lab for comments and critical reading of the manuscript. References [1] N.J. Pace, E. Weerapana, Diverse functional roles of reactive cysteines, ACS Chem. Biol. 8 (2013) 283–296. [2] R.K. Cannan, B.C. Knight, Dissociation constants of cystine, cysteine, thioglycollic acid and alpha-thiolactic acid, Biochem. J. 21 (1927) 1384–1390. [3] Z.R. Gan, W.W. Wells, Identification and reactivity of the catalytic site of pig liver thioltransferase, J. Biol. Chem. 262 (1987) 6704–6707. [4] Y.F. Yang, W.W. Wells, Identification and characterization of the functional amino acids at the active center of pig liver thioltransferase by site-directed mutagenesis, J. Biol. Chem. 266 (1991) 12759–12765. [5] M. Lo Conte, K.S. Carroll, The redox biochemistry of protein sulfenylation and sulfinylation, J. Biol. Chem. 288 (2013) 26480–26488. [6] N. Gould, P.T. Doulias, M. Tenopoulou, K. Raju, H. Ischiropoulos, Regulation of protein function and signaling by reversible cysteine S-nitrosylation, J. Biol. Chem. 288 (2013) 26473–26479.

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