Chemical Methods for Encoding and Decoding of Posttranslational Modifications

Please cite this article as: Chuh et al., Chemical Methods for Encoding and Decoding of Posttranslational Modifications, Cell Chemical Biology (2016), http://dx.doi.org/10.1016/j.chembiol.2015.11.006

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Review Chemical Methods for Encoding and Decoding of Posttranslational Modifications Kelly N. Chuh,1 Anna R. Batt,1 and Matthew R. Pratt1,2,* 1Department

of Chemistry of Molecular and Computational Biology University of Southern California, Los Angeles, CA 90089, USA *Correspondence: [email protected] http://dx.doi.org/10.1016/j.chembiol.2015.11.006 2Department

A large array of posttranslational modifications can dramatically change the properties of proteins and influence different aspects of their biological function such as enzymatic activity, binding interactions, and proteostasis. Despite the significant knowledge that has been gained about the function of posttranslational modifications using traditional biological techniques, the analysis of the site-specific effects of a particular modification, the identification of the full complement of modified proteins in the proteome, and the detection of new types of modifications remains challenging. Over the years, chemical methods have contributed significantly in both of these areas of research. This review highlights several posttranslational modifications where chemistry-based approaches have made significant contributions to our ability to both prepare homogeneously modified proteins and identify and characterize particular modifications in complex biological settings. As the number and chemical diversity of documented posttranslational modifications continues to rise, we believe that chemical strategies will be essential to advance the field in years to come. Introduction In the postgenomic era, it has become clear that the complexity of life cannot be explained by the number of genes in the genome alone. One layer of added functional and structural diversification beyond the genome is afforded via posttranslational modifications (PTMs). PTMs are covalent additions introduced to amino acid side chains or termini of proteins, either enzymatically or chemically, and represent one of the basic mechanisms to increase the chemical and biological diversity of the genome. These modifications range from the simple addition of a phosphate to the incorporation of large oligosaccharide structures, and they have been shown to change the biochemical and biophysical properties of the substrate protein. In addition to regulating activity, localization, and interactions with other proteins, PTMs can also carry information about the cellular environment (e.g., normal or disease state) or biochemical changes in response to various stimuli. PTMs can be dynamic in nature, and in many cases, cells are equipped with enzymatic machinery with opposing activities to install and remove the modification when given a functionally relevant cue. Despite the documented importance of PTMs in cellular biology, their identification and the study of specifically modified substrate proteins remain challenging. Although proteins can be harvested from cells for study, this process requires tedious and often difficult separation of their modified and unmodified forms. Furthermore, PTMs can occur on several sites simultaneously and substoichiometrically, making the isolation of a completely homogeneous population extremely difficult. Therefore, access to site-specifically modified proteins is of the utmost importance for the study of PTMs. In addition, identifying all proteins within the proteome that are substrates for a specific PTM continues to be a challenge despite being critical for understanding the biological pathways that control and are regulated by a given PTM. Unfortunately, some of the traditional tools for performing these types

of analysis (e.g., antibodies) are not available for all PTMs and cannot a priori distinguish enzyme-specific modification events. Over the years, many different approaches for studying PTMs have emerged, including the development of selective and unique chemical methods for the synthesis, identification, and analysis of posttranslationally modified proteins. Here, we review the methods that have been developed to encode and decode PTMs (Figure 1), where encode relates to the chemical synthesis or semisynthesis of homogeneously modified proteins or peptides, and decode defines the methods that are utilized for the isolation and identification of substrate proteins. This review focuses on modifications where chemical methods have been used to both encode and decode their function. For readers interested in PTMs that have only been addressed by one approach, we direct readers to other excellent reviews (Chuh and Pratt, 2015a; Davis and Chin, 2012; Grammel and Hang, 2013; Muir, 2003; Vila-Perello´ and Muir, 2010). Phosphorylation Protein phosphorylation is the transfer of an inorganic phosphate group to a variety of amino acid side chains, including most commonly to the hydroxyl groups of serine, threonine, and tyrosine residues (Figure 2A). The modification is installed by members of the kinase family of enzymes, which transfer the high-energy gamma phosphate from ATP to the substrate residues. Phosphorylation can be subsequently removed by phosphatase enzymes, rendering the modification dynamic. The first protein kinase, protein kinase A, was discovered in 1981 as the enzyme that could phosphorylate and subsequently activate the metabolic enzyme phosphorylase (Hayes and Mayer, 1981). This discovery would be just the tip of the iceberg, as protein phosphorylation networks have since been identified that control essentially all biological processes, including metabolic regulation, cellular growth and movement, and immune

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Review

Figure 1. Encoding and Decoding Posttranslational Modifications This review covers the different methods available in the chemical toolbox for either the preparation of site-specifically modified proteins (encoding) for subsequent biological experiments or the visualization and identification (decoding) of modifications from living systems and complex protein mixtures.

signaling. Therefore, it should not be surprising that altered phosphorylation plays critical roles in a variety of human diseases, including cancer (Gross et al., 2015; Zhang et al., 2009), neurodegeneration (Wang et al., 2004), and diabetes (Prada and Saad, 2013; Winder and Hardie, 1999). Accordingly, there is significant interest in targeting small-molecule inhibitors to kinases that are associated with specific pathologies. The first significant success in this area was the development of imatinib for the inhibition of the BCR-Abl kinase that drives chronic myelogenous leukemia (Capdeville et al., 2002). This has been followed by the clinical development of other inhibitors of growth factor receptor kinases, Bruton’s tyrosine kinase (Pan et al., 2007), and others. Despite these exciting results, gaining a complete understanding of the many roles of phosphorylation remains extremely challenging. There are approximately 500 kinases in the mammalian genome (Manning et al., 2002), 100 protein phosphatases that can antagonize these modifications (Alonso et al., 2004), and hundreds of proteins with phosphate-binding domains to read out a site-specific, phosphorylated signal (Yaffe, 2002; Yaffe and Elia, 2001). Complicating this system further, the precise timing and spatial localization of the modifications also play key roles in determining the cellular response to phosphorylation signals. Importantly, chemistry has made significant contributions to both the preparation and the analysis of site-specifically phosphorylated proteins and the development of new methodologies that link a specific kinase to its substrates. Encoding Phosphorylation The majority of posttranslationally modified proteins, including phosphorylated ones, have been generated in vitro using protein semisynthesis. Protein semisynthesis relies on the native chemical ligation (NCL) reaction for the preparation of full-length proteins from smaller fragments (Figure 2B) (Dawson et al., 1994). Since these smaller fragments are accessible by solid-phase peptide synthesis, they can be chemically modified with essentially any functionality, including PTMs and PTM analogs. NCL relies on the reversible transthioesterification reaction between

Figure 2. Encoding Protein Phosphorylation (A) A family of
500 kinases will transfer a phosphate group to certain amino acid side chains, including serine, threonine, tyrosine, and histidine. (B) Proteins can be synthesized using native chemical ligation (NCL). NCL involves the specific reaction of C-terminal thioesters and N-terminal cysteine residues to form native amide bonds. (C) Recombinant protein thioesters for use in NCL reactions can be created using proteins termed inteins, which catalyze the formation of a branched protein thioester that can be intercepted with exogenous thiols. (D) Unnatural amino acids can be site-specifically incorporated into proteins by using a combination of a tRNA synthetase enzyme that will charge an amber suppressor tRNA with an unnatural amino acid and a corresponding amber stop codon in mRNA.

a peptide/protein with a C-terminal thioester and another peptide/protein with an N-terminal cysteine residue, which results in a thioester linkage between the two fragments. This is quickly followed by an essentially irreversible sulfur to nitrogen acyltransfer, resulting in a native amide bond. This reaction proceeds in water, often with or without denaturants, and without the need for protecting groups on the amino acid side chains or PTMs of interest. Several solid-phase resins have been developed for the preparation of synthetic peptide thioesters using either Boc- or Fmoc-based peptide synthesis chemistries, and these peptides can then be reacted with either synthetic or recombinant protein fragments that have N-terminal cysteines. In the other orientation, recombinant protein thioesters can be prepared by taking advantage of the naturally occurring process of protein splicing (Figure 2C) (Vila-Perello´ and Muir, 2010). Specifically, proteins

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Review or protein fragments can be genetically fused in-frame to proteins termed inteins. These inteins will catalyze a nitrogen to sulfur acyl-transfer to generate a linked protein thioester that can be intercepted with exogenous thiols to give recombinant protein thioesters. These proteins can then undergo a variant of NCL, termed expressed protein ligation (EPL) (Muir et al., 1998). Notably, the recent development of ultra-fast inteins has greatly improved the efficiency of the preparation of protein thioesters compared with commercially available intein constructs (Shah et al., 2012; Shah and Muir, 2013). Phosphorylated tyrosine, serine, and threonine residues and their corresponding non-hydrolyzable mimics have been site specifically incorporated into proteins using semisynthesis. The first phosphorylated protein to be prepared using semisynthesis was the tyrosine kinase Csk, which was prepared bearing a phosphorylated tyrosine residue in its C terminus during the initial report of EPL (Muir et al., 1998). Csk is responsible for the phosphorylation of Src and Src family members at their C-terminal tail regions, which leads to a conformational change driven by an intramolecular interaction between the phosphorylated tail and SH2 domain in Src. Using EPL, the authors prepared an unnatural chimera between Csk and a C-terminal phosphorylated peptide and showed that this results in an intramolecular interaction that increases the kinase activity of Csk toward its substrates. Native phosphorylated serine and threonine residues have also been incorporated into proteins using semisynthesis. One of the best examples of this has been the series of experiments aimed at understanding the transforming growth factor b (TGF-b) signaling pathway (Massague´, 2012). Extracellular TGF-b will engage with TGF-b receptor kinase (TGF-bR), which will then go on to activate Smad transcription factors through the phosphorylation of two serine residues in their extreme C termini. During this process, a subunit of the receptor itself becomes phosphorylated on several serine residues. Using NCL, a soluble fragment of the TGF-bR was prepared with four such phosphorylation sites on its N terminus (Huse et al., 2001). In vitro studies with this protein demonstrated that it has improved binding to its substrate, Smad2, and reduced affinity for the protein inhibitor of the pathway, FKBP12. The consequences of the downstream phosphorylation on Smad2 were then examined. Using EPL, Smad2 was prepared bearing different phosphorylation patterns at serines 465 and 467, revealing that the trimerization of the transcription factor is largely driven by phosphorylation at residue 465, with a smaller contribution from modification of serine 467 (Ottesen et al., 2004). In a subsequent elegant display of the flexibility of EPL, phosphorylated Smad2 was also prepared with photoactivatable groups and fluorescent dyes, enabling the analysis of the kinetics of relocalization of trimerized Smad2 to the nucleus (Hahn and Muir, 2004; Pellois et al., 2004). In addition to the incorporation of natural phosphorylated residues, protein semisynthesis has also been used for the site-specific incorporation of non-hydrolyzable, difluoro- and non-substituted-methylene phosphonate analogs. In the case of tyrosine phosphorylation, these analogs have been most useful for the study of protein tyrosine phosphatases (PTPases). These enzymes can be phosphorylated at their C termini, but the effect of these modifications is difficult to study as the PTPase will remove their own phosphorylation marks. Incorporation of non-hydrolyzable analogs of

phosphorylated tyrosine using EPL overcame this limitation and showed that phosphate modifications had site-specific effects on the PTPase enzymatic activity (Lu et al., 2001, 2003; Schwarzer et al., 2006; Zhang et al., 2003). Serine and threonine phosphorylation have also been studied using the same approach. For example, EPL was used to prepare a semisynthetic version of casein kinase II alpha (CK2a) with a difluorophosphonate at threonine residue 344. A subsequent combination of microinjection and in vitro assays demonstrated that this modification increases the cellular stability of CK2a and alters its substrate selectivity (Tarrant et al., 2012). More recently, Lashuel and coworkers used EPL to prepare both unmodified and phosphorylated versions of the Huntingtin exon 1 protein that is prone to aggregation that causes Huntington’s disease (Ansaloni et al., 2014). Using these proteins, the author showed that phosphorylation at threonine residue 3 significantly slows the aggregation of this protein. Although semisynthesis has been quite powerful for the preparation of phosphorylated proteins, it does require some synthetic expertise and in some cases the refolding of proteins after purification. Another strategy that has the potential to overcome these issues, and has been used with success for the site-specific incorporation of phosphorylated residues or their structural analogs, is based on the use of the expanded genetic code and unnatural amino acids (Figure 2D). This technique relies on the recognition of a stop codon (typically the amber stop codon UAG) by an engineered tRNACUA that has been chemically or enzymatically charged with an unnatural amino acid, thereby generating an orthogonal codon that can be read out by the ribosome during protein translation (Chin, 2014; Lang and Chin, 2014; Liu and Schultz, 2010). Dozens of unnatural amino acids have now been incorporated into proteins in Escherichia coli, yeast, and mammalian cells using this system. One of the most successful orthogonal aminoacyl-tRNA synthetase/tRNACUA pairs has been based on the tyrosine pathway (TyrRS/tRNATyr) from Methanococcus jannaschii. Although this system has not yet been engineered to incorporate phosphorylated tyrosine, it has been used to site specifically incorporate the tyrosine analog, p-methoxylmethyl-phenylalanine, into recombinant protein in E. coli (Xie et al., 2007). Specifically, the target protein this study focused on was the transcription factor signal transducer and activator of transcription-1 (STAT1), and the procedure generated a constitutively active protein. The approach developed to incorporate phosphorylated serine into proteins is somewhat different and takes advantage of an interesting twostep pathway for the introduction of cysteine residues into proteins used by some methanogenic archaea. For these organisms, the first step in making cysteine aminoacylated tRNA needed for protein synthesis, is to attach phosphorylated serine onto a tRNA (tRNACysGCA) using the phosphoseryl-tRNA synthetase (SepRS). Then a second enzyme converts the phosphorylated serine to cysteine to give tRNACysGCA that is ready for protein synthesis. This raises an interesting question of how methanogenic archea select for the final product, tRNACysGCA, and avoid incorporating the intermediate tRNA carrying a phosphorylated serine. One explanation put forward was that the corresponding tRNASepGCA would be a poor substrate for the elongation factor EF-Tu and thus discriminated against by the protein synthesis machinery. This suggested that EF-Tu

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Cell Chemical Biology

Review Figure 3. Decoding Protein Phosphorylation (A) Development of analog-sensitive kinases using a bump-hole strategy. Wild-type kinases are incapable of using the bumped ATP analog N6benzyl ATP. However, mutation of the kinase in its active site creates a hole that will allow N6-benzyl ATP to function as a substrate. (B) Identification of kinase substrates using analog-sensitive kinases. A gatekeeper mutant kinase of interest is first incubated with cell lysate and N6-benzyl-ATPɣS, resulting in selective thiophosphorylation of that kinase’s substrates. The resulting thiophosphate is then alkylated to generate a p-nitro-benzyl group that is recognized by a specific antibody for visualization or enrichment. (C) Linking a known phosphorylated substrate with the kinase responsible using cross-linking. An ATP based cross-linker is first incubated with a complex mixture of kinases in a cell lysate, transferring a Michael acceptor to the conserved, catalytic lysine residue. Then a substrate peptide bearing a cysteine residue at the known site of phosphorylation is added, yielding a covalent cross-link between the substrate and kinase of interest.

could be engineered to favor incorporation of phosphorylated serine into recombinant proteins. This was done and worked as expected (Lee et al., 2013; Park et al., 2011) although the yields were lower than for other unnatural amino acid systems. More recently, this same orthogonal system was improved to allow for the incorporation of phosphorylated serine and its difluoro-phosphonate analog (Rogerson et al., 2015). Specifically, engineering of the aminoacyl-tRNA synthetase/tRNACUA pair (SepRS/tRNApSerCUA) yielded mutants that will incorporate phosphorylated serine into recombinant proteins in E. coli 18 times more efficiently without the need for engineering of EFTu. Notably, in E. coli, phosphorylated serine is generated in the serine biosynthesis pathway and this endogenous reaction can serve as the source of modified serine by the system. In addition, mutation of this biosynthetic pathway to reduce the cellular concentration of phosphorylated serine enabled the incorporation of an exogenously added phosphonate analog of serine. Undoubtedly, the further optimization of these technologies, including other orthogonal synthetase/tRNA pairs, will allow for the incorporation of both phosphorylated tyrosine and serine in a variety of cell types. Decoding Phosphorylation The identification of phosphorylated proteins under different cellular states in different tissues, etc. is critical for understanding the specific roles of this PTM. Site- and pan-specific antibodies can give a broad picture of the level of individual or global phosphorylation levels, respectively, and enrichment methods (e.g., metal affinity chromatography) coupled with proteomics have enabled the global characterization of the phosphopro-

teome. However, mapping kinases to their specific substrates has been more difficult and has inspired the creation of different chemical methods. The first such method uses engineered protein kinases, termed analog-sensitive kinases, that are optimized to exclusively accept an analog of ATP that cannot be utilized by any wild-type kinases (Figure 3A) (Liu et al., 1998; Shah et al., 1997; Ubersax et al., 2003; Zhang et al., 2005). Specifically, this method takes advantage of the fact that most kinases have a large (threonine, methionine, etc.) conserved residue in the ATP-binding pocket. This gatekeeper residue prevents the binding of an N6-benzyl-modified ATP in the kinase active site. However, when the gatekeeper residue is mutated to a smaller amino acid (e.g., glycine), the resulting analog-sensitive kinases will use bulky ATP derivatives, such as N6-(benzyl) ATP, to phosphorylate their substrates. This method has been generally termed the ‘‘bump-hole’’ strategy. When radioactive N6-benzyl ATP is used with a specific analog-sensitive kinase, the substrates of that kinase can be visualized in a complex cellular lysate. Notably, these same mutant kinases can be selectively inhibited by bulky ATPcompetitive kinase inhibitors (Bishop et al., 1998, 2000). By genetically incorporating the mutant kinase into a cell or organism, one can achieve highly selective inhibition and investigate the phenotypic and biochemical effects. While this method enables the visualization of specific kinase substrates and has been shown to be broadly applicable, it does not immediately allow for the unbiased identification of these substrates from a complex mixture. To accomplish this, the fact that kinases will transfer the ATPgS analog to thiophosphorylate proteins was exploited (Allen et al., 2005, 2007). In this elegant strategy, the resulting protein thiophosphate can then be alkylated with a p-nitro-benzyl group (Figure 3B). An antibody that specifically recognizes this alkylated structure can then be used to either visualize or enrich kinase substrates for proteomic identification.

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Review Combining analog-sensitive kinases with this approach allows, in principle, for the enrichment and identification of the substrates of any kinase of interest. Subsequent iterations of this strategy have used direct alkylation of the thiophosphate by solid-phase resins followed by their selective elution (Blethrow et al., 2008) and the selective capping of cysteine residues to eliminate the potential enrichment of false-positive proteins (Garber and Carlson, 2013). The thiophosphate method above starts with a kinase of interest and proceeds to the identification of the corresponding substrate proteins. An equally important challenge is to map a known phosphorylated protein back to the kinase responsible for the modification. Again chemistry contributed to solving this problem, in particular with the development of several small-molecule cross-linkers. One class of these compounds relies on a three-component reaction between a substrate protein (or peptide) of interest, the small molecule, and the conserved active-site lysine residue of a kinase (Maly et al., 2004; RielMehan and Shokat, 2014; Statsuk et al., 2008). All of these cross-linkers rely on the genetic incorporation of a cysteine at the normal site of phosphorylation in the substrate. In the most robust system to date, the cross-linker is an analog of ATP that binds to the kinase active site and results in the modification of the catalytic lysine to generate a methacrylamide, which will then undergo a Michael-addition reaction with the cysteine-containing substrate bait (Figure 3C). This method has been successfully applied to a model system in a complex lysate, but unfortunately, not to date for the unbiased identification of a kinase-substrate pair. An alternative strategy was also explored that relied on the incorporation of two photo-cross-linking groups into an analog of ATP (Parang et al., 2002). Upon photo-irradiation, one of the cross-linkers near the adenosine ring covalently labels the kinase, while the other located near the gamma phosphate traps the substrate protein. This system was successfully applied to an in vitro model system but has not been employed in a large-scale discovery experiment. Glycosylation Glycosylation is a common PTM where carbohydrate chains of various lengths and composition are added to a large fraction of proteins, highlighting the utility of glycosylation in biological events. Specifically, proteins that reside in the secretory pathway, at the cell surface, or are excreted into extracellular space, can be modified by typically large and elaborate oligosaccharides in the ER and Golgi, resulting in N-linked and mucin O-linked glycosylation. The most common type of O-linked glycosylation, mucin-type glycosylation, is characterized by the core addition of N-acetyl-galactosamine (GalNAc) through an a-O-linkage to the b-hydroxyl group of serine or threonine residues. This monosaccharide can then be elaborated through addition of sialic acid, fucose, and/or additional units of Galb1,4GlcNAc to form large, branched glycan structures. Mucin-type glycosylation (Hang and Bertozzi, 2005) is found on many cell-surface proteins and has been shown to play an essential role in protein localization and cell-cell communication in the immune response (Wolfert and Boons, 2013). A more prevalent type of protein glycosylation is N-linked glycosylation. In contrast to the synthesis of mucin-type glycoproteins, N-linked glycoproteins are synthesized first by the assembly of a doli-

chol-linked oligosaccharide precursor in the cytosol and ER. The large structure is subsequently transferred by an oligosaccharide transferase to the amide side chain of an asparagine residue of a nascent polypeptide. Intracellularly, N-linked glycans regulate protein trafficking and act as quality control for protein folding (Helenius and Aebi, 2001). Outside the cell, N-linked glycans can function as ligand receptors and have been shown to mediate cell interactions with proteins, other cells, and pathogens. Serine and threonine residues of intracellular proteins can also be O-glycosylated by the single monosaccharide, N-acetyl-glucosamine (GlcNAc). Termed, O-GlcNAc, this PTM is known to affect protein localization and signal transduction. In fact, altered states of O-GlcNAc glycosylation have been associated with oncogenic transformation, neurodegenerative disease, and perhaps diabetes (Dias and Hart, 2007; Ma and Vosseller, 2013). All types of glycoproteins are fundamental in biology and therefore have conjured a great deal of interest within the scientific community. Their study, however, has proved challenging due to the fact that naturally occurring glycoproteins are not synthesized homogeneously and current chromatographic methods are not sophisticated enough to separate the variety of glycoforms on a reasonable scale. Furthermore, complex oligosaccharide structures are not synthesized in a template-dependent manner; therefore, no straightforward genetic methods for controlling expression of specific carbohydrates exist, leaving researchers with a restricted set of tools for their study. Therefore, there is a demand for a source of structurally homogeneous glycoproteins for functional studies (Davis, 2002; Gamblin et al., 2009; Grogan et al., 2002), which involves methodologies from carbohydrate and peptide chemistries alike, as well as the creation of chemical tools for the visualization and identification of glycoproteins. Encoding Glycosylation The most common method to synthesize glycopeptides/glycoproteins relies heavily on solid-phase peptide synthesis (SPPS) to incorporate glycosylated amino acids onto growing amino acid chains resulting in a native glycan-amino acid bond. The majority of naturally occurring, O-linked glycans can be synthesized by the glycosylation of protected serine or threonine residues using common glycosyl donors, resulting in a-O-linked cassettes for SSPS (Figure 4A). For example, the Boons laboratory has demonstrated the synthesis of the Tn and Tf antigen building blocks using the thiophilic Ph2SO/Tf2O promoter system for subsequent glycosylation of an Fmoc-protected threonine for the synthesis of glycopeptides (Cato et al., 2005). For the synthesis of larger glycan structures, the core a-O-Ser/Thr linkage is formed first, and orthogonally removable protecting groups can then be used to direct the additional elaboration of branching sugars. Impressively, this linear, cassette-based method has been employed by the Kunz laboratory in the synthesis of part of the CD62P ligand PSGL-1 that is involved in the inflammatory response and leukocyte recognition. The resulting 18-residue fragment containing an O-linked hexasaccharide was synthesized in sufficient amounts for use in biomedical studies (Baumann et al., 2008). In the synthesis of N-linked glycopeptides, glycosylated asparagine cassettes are most commonly prepared before or after SPPS through the reaction of a protected or deprotected anomeric glycosyl-amine with an aspartate residue (Figure 4B). For example, the Keissling

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Review

Figure 4. Protein Glycosylation (A) O-linked glycopeptides are typically synthesized through the solution-phase preparation of an Fmoc-protected amino acid cassette that can be used directly in solid-phase peptide synthesis. (B) N-linked glycopeptides have been prepared using the cassette approach but can alternatively be synthesized after peptide synthesis through the coupling of an aspartic acid residue to a glycosyl-amine or amine equivalent. (C) Enzymatic installation of large N-linked glycans onto peptides and proteins with transglycosylation reactions. Under certain reaction conditions, some endoglycosidases will use isolated or synthesized glycans as substrates and transfer them onto single N-acetyl-glucosamine residues on peptides or proteins. (D) Metabolic chemical reporters of glycosylation. Living cells are treated with analogs of monosaccharides containing bioorthogonal functionality (e.g., an alkyne). These reporters are metabolized by the cell and installed onto proteins. Bioorthogonal reactions can then be performed for the installation of visualization or affinity tags. (E) Chemoenzymatic detection of O-GlcNAc modifications. Endogenous O-GlcNAc modifications in a cell lysate can be enzymatically modified with a GalNAz residue, followed by the installation of tags using bioorthogonal chemistry.

laboratory has developed a method in which the glycosyl-amide bond is formed through reaction with glycosyl azides and asparagine-derived phosphinothioesters, avoiding anomerization and formation of isomeric mixtures (He et al., 2004). More recently, Doores and coworkers have demonstrated an elegant procedure for glycosyl-asparagine ligation that can be utilized with fully deprotected substrates, is stereocontrolled, and is compatible with linear and convergent approaches that are free from the need of complex auxiliaries (Doores et al., 2006). The development of site-selective chemical glycosylation of proteins has allowed for the synthesis of larger and more complex glycopeptides; however, site-specific installation of native glycans in full-length proteins remains challenging. To overcome the size limitation of SPPS, larger glycoproteins can be prepared in a stepwise fashion using NCL. For example, to mechanistically explore the role of N-linked glycosylation during protein folding in the ER, Izumi et al. (2012) synthesized misfolded interleukin-8 (IL-8,

CXCL8) bearing an N-linked glycan. Using this material, they found that the enzyme UDP-glucose:glycoprotein glucosyltransferase (UGGT) prefers misfolded over correctly folded glycoproteins, indicating that UGGT plays a distinct role in quality control of protein folding. Extending the use of NCL, the Danishefsky group published the complete chemical synthesis of the signaling glycoprotein, erythropoietin, with all carbohydrate domains at all native glycosylation sites (Wang et al., 2013a). Our laboratory has utilized EPL to study the function of O-GlcNAc modification on the protein a-synuclein, the major aggregating protein associated with Parkinson’s disease and dementia with Lewy Bodies. Chemical synthesis of homogeneous, full-length, O-GlcNAcylated a-synuclein was achieved using EPL in three steps, and subsequent aggregation assays showed that O-GlcNAc modification at threonine residue 72 inhibits a-synuclein aggregation and is non-toxic to neurons in cell culture (Marotta et al., 2015).

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Review Extension of these synthetic techniques to larger glycans has been stymied due to the low efficiency of couplings and the complicated nature of both the carbohydrate and amino acid protecting group chemistries involved in SPPS. In addition, some glycosidic bonds cannot survive the strong acidity of final deprotection conditions of peptides (e.g., trifluoroacetic acid [TFA]). To address these shortcomings, chemoenzymatic methods to catalyze the addition of single monosaccharides onto synthetic glycopeptides or growing glycan structures have been developed as an alternative. Sometimes referred to as glycoprotein remodeling, recombinant enzymes with unique glycosyltransferase/glycosidase activity can be utilized in a stepwise fashion to produce more complex glycopeptide libraries that are out of reach of canonical chemical synthesis. For example, Bello et al. (2014) recently demonstrated the stepwise chemoenzymatic synthesis of O-linked glycosylated MUC1 peptides utilizing Drosophila glycosyltransferases. In the area of N-linked glycoproteins, the Boons and Paulson laboratories collaborated to create a library of complex multi-antennary glycans by utilizing a chemically synthesized core oligosaccharide to which asymmetric branched antenna were chemoenzymatically installed (Wang et al., 2013c). The resulting library was printed on a microarray and was subsequently screened for binding to lectins and influenza-virus hemagglutinins (HAs). The results illustrate the complex environment-dependent recognition of glycan epitopes and highlight the importance of understanding the receptor specificity to further elucidate their biological consequences in disease. Incorporating complex N-glycans into peptides can also be difficult due to the involved chemistry required for their synthesis and conjugation to aspartic acid. In order to allow for the use of isolated N-linked glycans and improve the coupling of isolated and synthesized structures onto peptides, endo-b-N-acetylglucosaminidases (ENGases) have been utilized to perform transglycosylation reactions (Figure 4C) (Wang, 2011). These enzymes normally cleave N-linked glycans, leaving only the first GlcNAc residue at the N-linked site(s). However, through mutagenesis and screening of reaction conditions, it was found that certain ENGases will essentially perform this reaction in reverse to install complex N-linked glycans onto single GlcNAc residues on peptides. In an early example of this method, the enzyme EndoM was used to prepare glycosylated versions of the N-linked glycopeptide hormone calcitonin, a 32-amino acid calcium-regulating hormone used in the treatment for hypercalcemia, Paget’s disease, and osteoporosis (Haneda et al., 1998). Other Endo enzymes have also been explored, such as EndoF2 and F3, that were found to glycosylate a-1,6-fucosylated GlcNAc derivatives, producing native, corefucosylated, complex-type glycopeptides (Huang et al., 2011). More recently, the Davis laboratory has explored a bacterial endoglycosidase, EndoS, that is complimentary to other endoglycosidases, EndoA and EndoH (Goodfellow et al., 2012). Specifically, the authors showed that a synthesized tetrasaccharide oxazoline could be transferred onto human IgG using EndoS, and that EndoS also shows tolerance to the presence of core fucosylation, broadening its synthetic utility. Although useful, chemoenyzmatic methods do not always allow for control over the site of glycosylation. Therefore, to further increase the selectivity of protein glycosylation, various chemoselective methods have been developed. Specifically,

preformed glycans are attached to peptides or proteins through mild and selective chemical reactions that tolerate numerous functional groups, therefore minimizing the need for protecting groups. The Bertozzi group pioneered this area in the synthesis of O-linked glycopeptides that contain unnatural bonds at the C-6 and C-3 branchpoints, as oximes (Rodriguez et al., 1997) and thioethers, respectively (Marcaurelle and Bertozzi, 2001), as well as thioether linkages to install antennae onto an N-linked core (Pratt and Bertozzi, 2003). More recently, other methods that take advantage of the native chemistry of the cysteine thiol have also been utilized. Notably, Dondoni et al. (2009) demonstrated a ligation strategy that utilizes the thiol-ene coupling (TEC) reaction in which an alkenyl C-glycoside is coupled via photo-irradiation with a protein or peptide containing a free sulfhydryl group resulting in a thioether bond. Thioglycosides are an attractive synthetic mimic due to their likeness in length to the native glycosidic bond as well as their increased stability (Chalker et al., 2011). The Davis group published a tag-andmodify method that involved the use of TEC chemistry in the synthesis of S-glycosyl amino acids through the addition of glycosyl-thiols to homoallylglycine (Hag) following its incorporation into a peptide/protein as a non-natural amino acid (Floyd et al., 2009). The tag-and-modify method has also utilized bioorthogonal azide-alkyne cycloaddition reactions. Specifically, through the introduction of the unnatural amino acid azidohomoalanine (Aha), GlcNAc-modified protein Np276 from Nostoc punctiforme was synthesized (Ferna´ndez-Gonza´lez et al., 2010). In a similar fashion, an alkyne-containing non-natural amino acid, homopropargylglycine (Hpg) can be incorporated as a methionine surrogate. Interestingly, the Davis laboratory utilized both Aha and Hpg to enable attachment of multiple glycans to bacterially expressed protein scaffolds (van Kasteren et al., 2007). Decoding Protein Glycosylation Different cellular states and tissues can display unique glycoproteins and glycan structures, and direct identification of modified proteins is essential to uncovering the role of this posttranslational modification. Pioneered by the Bertozzi laboratory, several laboratories have investigated the use of metabolic chemical reporters (MCRs) for the visualization and identification of glycosylated proteins (Figure 4D). Typically, this technique involves treatment of cells with chemically synthesized analogs of naturally occurring monosaccharides that contain bioorthogonal reactivity (commonly an azide or alkyne). These MCRs are then accepted by living systems and metabolically converted into high-energy UDP-sugar donors that are subsequently utilized in their incorporation into glycans by endogenous glycosyltransferases. The first completely orthogonal MCR, Ac4ManNAz, was developed by the Bertozzi laboratory (Saxon and Bertozzi, 2000). Once this compound diffuses into cells, the O-acetates are removed by endogenous lipases and the resulting ManNAz is biosynthetically converted to the corresponding azide-containing sialic acid analog and enzymatically installed onto the termini of cell-surface glycans. Subsequent bioorthogonal labeling of the azide, using reactions like the Staudinger ligation or the copper-catalyzed azide-alkyne cycloaddition, can then be used for the selective installation of visualization or affinity tags. Since this transformative initial study, many different MCRs of glycosylation have been developed, including other azide-bearing

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Cell Chemical Biology

Review monosaccharides like Ac4GalNAz (Hang et al., 2003), Ac4GlcNAz (Vocadlo et al., 2003), and Ac4FucAz for the labeling of mucin O-linked glycans, O-GlcNAc modifications, and fucose-containing glycans, respectively. Our laboratory subsequently found that the alkyne MCR, Ac4GlcNAlk, displayed improved signal to noise and was more selective for O-GlcNAc modifications compared with Ac4GlcNAz (Zaro et al., 2011), and other alkyne MCRs for the visualization of sialic acid and fucose have also been developed (Hsu et al., 2007). Furthermore, we have continued to make structural changes to these MCRs, resulting in the recent discovery of an MCR, Ac36AzGlcNAc, that is selective for O-GlcNAc modifications (Chuh et al., 2014), to our knowledge the first and only glycosylation-type specific MCR. Currently, there are also monosaccharide MCRs that contain other bioorthogonal functionalities, including alkenes (Niederwieser et al., 2013; Spa¨te et al., 2014) and cyclopropenes (Cole et al., 2013; Patterson et al., 2012, 2014), which can take advantage of the rapid tetrazine ligation for the installation of tags. Notably, monosaccharide MCRs have been used in a variety of contexts from cell culture to living animals for the visualization (Chuh and Pratt, 2015b) and proteomic identification (Chuh and Pratt, 2015a) of glycoproteins. For example, Ac4ManNAz and Ac4GalNAz have been used to visualize cell-surface glycans in zebrafish embryos (Baskin et al., 2010; Laughlin et al., 2008) and Caenorhabditis elegans (Laughlin and Bertozzi, 2009). In addition, many monosaccharide MCRs have been used to perform proteomic analysis of different glycoproteins, including quantitative comparisons of cancer versus normal cell populations using Ac4GalNAz (Slade et al., 2012). Recently, the Bertozzi laboratory has developed a technique termed isotope-targeted glycoproteomics (IsoTaG), a mass-independent chemical glycoproteomics method for the identification of intact metabolically labeled (using Ac4ManNAz or Ac4GalNAz) glycopeptides from the total proteome (Woo et al., 2015). In contrast to traditional tandem mass spectrometry (MS/MS) proteomics that is performed on the most abundant species in the total-scan mass spectra, IsoTaG enables the specific detection of glycoproteins with isotopic signatures, improving the selection of low-abundance glycopeptides. MCRs have been transformative in their ability to report on different types of glycoproteins; however they necessarily compete with natural metabolites, meaning that they are inherently unreliable indicators of the overall levels of a modification. To address this limitation in the area of O-GlcNAc modification, the Hsieh-Wilson laboratory has developed chemoenzymatic detection methods that enable the capture of a snapshot of endogenous O-GlcNAc-modified proteins. In the most common iteration of this technology, incubation of cell lysates with a recombinantly expressed, mutant b-1,4-galactosyltransferase and chemically prepared UDP-GalNAz results in the transfer of GalNAz to O-GlcNAc residues (Figure 4E) (Clark et al., 2008). The resulting azide-containing disaccharide can be bioorthogonally reacted with different visualization and affinity tags. For example, small polyethylene glycol (PEG) chains can be installed that will shift the mass of O-GlcNAc-modified proteins, enabling the stoichiometry of some O-GlcNAc modifications to be quantitated using western blotting (Ortiz-Meoz et al., 2014; Rexach et al., 2010). In addition, this strategy has been combined with both b-elimination (Khidekel et al., 2007) and electron transfer dissociation MS (e.g., Alfaro et al., 2012)

for the proteomic identification of O-GlcNAcylated proteins and a subset of modification sites. Ubiquitination The covalent addition of a small (76-residue) protein called ubiquitin to lysine side chains of protein through isopeptide bonds is a posttranslational modification that is implicated in numerous cellular processes including proteasomal degradation, signal transduction, receptor endocytosis, and DNA damage response (Figure 5A) (Chen and Sun, 2009; Komander and Rape, 2012). Ubiquitin is added by an enzymatic cascade that involves three classes of proteins: E1 ubiquitin-activating enzymes, E2 ubiquitin-conjugating enzymes, and E3 ubiquitin ligases (Hershko and Ciechanover, 1998). First, the C-terminal glycine residue of ubiquitin is activated by ATP to yield a thioester intermediate with the catalytic cysteine of the E1 enzyme. Then, ubiquitin is transferred onto an E2 conjugating enzyme to generate a second active-site thioester intermediate. The final step of the ubiquitin enzymatic cascade is catalyzed by an E3 ligase to form an isopeptide bond between the C-terminal glycine reside of ubiquitin and the ε-amino group of the target lysine reside of the substrate protein. Interestingly, proteins can be modified with either a single ubiquitin molecule (monoubiquitination) or ubiquitin chains (polyubiquitination), both of which give rise to a plethora of ubiquitin signals. Specifically, monoubiquitination can be found at more than one site, giving rise to multi-monoubiquitination. On the other hand, polyubiquitination can be formed through one of the seven ubiquitin lysine resides (Lys6, Lys11, Lys27, Lys29, Lys33, Lys48, and Lys63) or through the amino terminal methionine residue to generate linear chains (Kulathu and Komander, 2012). Given all the possible outcomes of ubiquitination, it is not surprising that this PTM accounts for a wide range of functions within the cell. Akin to phosphorylation, ubiquitination is a dynamic modification and can be removed from protein substrates by the action of the enzyme family of deubiquitinases (DUBs) (Komander et al., 2009). Given its pervasive role in cellular function, understanding the role of ubiquitination is of the utmost importance, and chemistry has contributed to this goal through both the synthesis of site-specifically ubiquitinated proteins and the development of probes for the identification of ubiquitin modifying enzymes. Encoding Ubiquitination Although the chemical synthesis of ubiquitin was achieved over 20 years ago through SPPS, the synthesis of ubiquitinated substrate proteins remained challenging until the advent of NCL. Muir and coworkers were the first to synthesize a monoubiquitinated protein, histone H2B, containing a native isopeptide linkage using EPL to study the mechanistic role of ubiquitination in enhancing subsequent methylation of lysine residue 79 in histone H3 (H3K79) (Figure 5B) (McGinty et al., 2008). The synthetic portion of the protein was a peptide corresponding to residues 117–125 of H2B. This peptide contained an N-terminal cysteine residue protected with a photo-labile o-nitrobenzyl group and a photo-labile, thiol-bearing ligation auxiliary attached to the ε-amino group of Lys120 through a glycine linker (eventually to become Gly76 of ubiquitin). This ligation auxiliary then mediated ubiquitination through an NCL reaction with a recombinant ubiquitin(1–75) C-terminal thioester, and the ligation auxiliary and the cysteine protecting group were subsequently removed

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Figure 5. Ubiquitination (A) Ubiquitination is the addition of the small protein ubiquitin to protein side chains, most often lysine, resulting in an isopeptide bond. This first modification event can then be polymerized in various ways to form polyubiquitin chains. (B) Synthesis of ubiquitinated histone H2B using a photo-cleavable auxiliary. Using an NCL reaction, ubiquitin is first installed onto a synthetic peptide through a lysine residue bearing the auxiliary. Photolysis then both removes the auxiliary and reveals the N-terminal cysteine residue that can be used in subsequent NCL reactions. (C) Ubiquitination of proteins using a d-mercapto-lysine residue. The d-mercapto-lysine residue is first incorporated into a peptide using solid-phase peptide synthesis, where it can then undergo an NLC reaction with a ubiquitin thioester. The d-thiol group is then removed by chemical desulfurization. (D) A d-mercapto-lysine residue can be site specifically installed into recombinant proteins using unnatural amino acid mutagenesis. (E) Examples of isopeptide linkages that have been used for the installation of ubiquitin.

by irradiation with UV light. A second EPL reaction was preformed between this ubiquitinated peptide and a recombinant H2B(1–116) C-terminal thioester to give full-length ubiquitinated H2B. Since H2B has no native cysteine residues, chemical desulfurization was used to convert the cysteine required for the ligation to alanine to yield the monoubiquitinated H2B protein with no mutations. This synthetic protein was used to show that monoubiquitination of H2B directly stimulates methylation at H3K79 by the methyltransferase hDot1L, and it inspired a series of important innovations in the synthesis of site-specifically ubiquitinated proteins. The first significant advanced was the synthesis of d-mercaptolysine unnatural amino acid building blocks for SPPS first developed by the Brik laboratory (Ajish Kumar et al., 2009; Kumar et al., 2010) followed shortly by Ovaa and coworkers (Figure 5C) (Oualid et al., 2010). Like the previous photo-liable ligation auxiliary, this amino acid can be directly incorporated into peptides and then undergo NCL reactions with ubiquitin C-terminal thioesters, followed by desulfurization for the site-specific installation of ubiquitin. When combined with heroic protein chemistry efforts, these amino acids have enabled the synthesis of quite large proteins. For example, the Parkinson’s disease associated protein a-synuclein was prepared bearing either mono-, di-, or tetraubiquitination at the physiologically relevant lysine residue 12, and the different effects on protein aggregation and stability were measured (Haj-Yahya et al., 2013). However, the size of many proteins can make their chemical synthesis difficult. To address this limitation, the Chin and Kommander laboratories collaborated to use unnatural amino acid mutagenesis to introduce their GOPAL (genetically encoded orthogonal protection and activated ligation) strategy (Virdee et al., 2010). Briefly, a lysine residue of interest can be genetically replaced with Nε-(tert-butyloxycaronyl)-L-lysine (NHBoc-Lys) using unnatural amino acid muta-

genesis and the Methanosarcina barkeri MS pyrrolysine tRNA synthetase (MbPyrlRS) and its corresponding amber suppressor tRNA (MbtRNACUA). After recombinant expression, the remaining lysine residues can be orthogonally protected by treatment with N-(benzyloxycarbonyloxy)succinimide (Cbz-OSu). After deprotection of the NHBoc-Lys, a suitably protected ubiquitin molecule can be site specifically installed, followed by global deprotection. Unfortunately, the GOPAL system relies on extensive protection group chemistry that can contribute to poor yields. To overcome this issue, the Chin laboratory designed a synthetic scheme in which a d-thiol-L-lysine is incorporated at the desired site of ubiquitylation using an improved pyrrolysyltRNA synthetase/tRNACUA pair (Figure 5D) (Virdee et al., 2011). Using this method, Chin and coworkers prepared K6-linked diubiquitin and site specifically ubiquitinated SUMO (small ubiquitin-like modifier protein) at K11 through the formation of native isopeptide bonds. A great deal of research has been dedicated to the synthesis of native isopeptide-bond conjugated ubiquitin through NCL and EPL and has allowed insight into the biological roles that this PTM can play. However, many of these approaches are synthetically challenging and require multiple ligation and purification steps, thus limiting the widespread use of NCL and EPL for the semisynthesis of ubiquitinated proteins. To address this shortcoming, several groups have created strategies to introduce ubiquitin onto substrate proteins through isopeptide bond mimics (Figure 5E). One of the first such methods was a disulfide-directed approach that was developed simultaneously by the Muir and Zhuang laboratories (Chatterjee et al., 2010; Chen et al., 2010). This approach takes advantage of a ubiquitin-intein fusion that is trapped with cysteamine to incorporate a thiol moiety at the C terminus of ubiquitin (Ub-SH), which can be subsequently activated as a mixed disulfide by treatment with specific

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Review reagents like 2,20 -dithiobis(5-nitropyridine) (DTNP). The target protein carrying a cysteine residue at the desired modification site is then reacted with the activated ubiquitin resulting in the formation of a disulfide between the substrate and ubiquitin moieties. Although this technique has been successfully applied to a handful of ubiquitination sites in vitro (e.g., Abeywardana et al., 2013; Meier et al., 2012), the disulfide linkage is not chemically stable. Therefore, other chemical approaches have been used for the installation of stable analogs of the ubiquitin linkage. For example, the Strieter laboratory took advantage of TEC to install thioether linkages for conjugation of ubiquitin to substrate proteins (Valkevich et al., 2012). Furthermore, they were able to show that this Nε-Gly-L-homothialysine isopeptide linkage was hydrolyzed by DUBs in a manner similar to that of the wildtype isopeptide bond, demonstrating that it is a good structural mimic. In yet another thioether approach, the Brik laboratory utilized either an a-bromo acetamide or maleimide synthetically tethered to the C terminus of ubiquitin that can be subsequently reacted with a cysteine of a substrate protein (Hemantha et al., 2014). Copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC) has also been used for the installation of ubiquitin to a substrate protein via a triazole linkage. For example, the installation of a cysteine into the target protein followed by treatment with iodoacetamide ethyl azide allows for the installation of a site-specific azide, which can be reacted with a ubiquitin bearing a C-terminal alkyne (installed using intein chemistry) (Weikart and Mootz, 2010). More recently, similar strategies have been developed that take advantage of the incorporation of azide- or alkynecontaining amino acids (Eger et al., 2011; Ro¨sner et al., 2015; Sommer et al., 2011). Additional methods based on the use of non-natural linkages continue to be developed and applied, including propanone (Yin et al., 2000) and oxime (Shanmugham et al., 2010) linkages. Decoding Protein Ubiquitylation Visualization and identification of ubiquitin enzymatic machinery has been accomplished using activity-based protein profiling (ABPP), which utilizes enzyme-specific probes that react, in most cases covalently, within the active site of various enzymes (Nomura et al., 2010). Although the targets of activity-based probes vary, typically they contain two elements: a reactive group containing an electrophile to react with a nucleophilic residue within the enzyme, and a tag. For example, a panel of radioactive- or HA-tagged, DUB-specific probes containing a C-terminal thiol-reactive group or warhead was synthesized using EPL. They were subsequently utilized as suicide substrates to first visualize and then identify active DUBs from mammalian cells (Borodovsky et al., 2001, 2002). The range of electrophilic warheads was then expanded beyond the vinylmethylester and vinylethoxysulfone groups used previously to enable the identification of both DUBs and E3 ligases (Love et al., 2009). These probes used known cysteine-selective electrophiles; however, an active-site directed probe containing an alkyne as the C-terminal electrophile was recently demonstrated to also react with active-site cysteines (Ekkebus et al., 2013). The researchers were surprised to discover that their C-terminally propargylated Ub (Ub-Prg), originally synthesized for site-specific ubiquitination of peptides, inhibited the human DUB ubiquitin C-terminal hydrolase isoenzyme L3 (UCHL3). Confirmed by X-ray crystallography, it was found that the resulting quaternary vinyl thio-

ether conferred selectivity toward de-ubiquitinating enzymes. Continued success in this field with these types of atypical warheads will undoubtedly enable the preparation of ever more selective probes and potentially extend this technique to the metalloprotease members of the DUB family. Lipidation The attachment of long-chain fatty acids to proteins, termed lipidation, is a PTM that regulates membrane affinity, localization, and trafficking (Figure 6A) (Hang and Linder, 2011). There are many types of lipid modifications and their covalent attachment to proteins has revealed a complicated network of membranes and lipidated proteins that are at the center of basic cellular function and human disease. This complexity, combined with an almost complete lack of appropriate biological reagents (e.g., antibodies), have increased the pressure to develop specific and sensitive methods to probe their function and identify modified substrates. Myristoylation and palmitoylation are the two most common classes of fatty-acylation events that typically occur co- and posttranslationally, respectively. Myristoylation is characterized by the irreversible covalent attachment of a 14-carbon fatty acid, myristic acid, to the N terminus of a substrate protein via an amide linkage that only occurs in eukaryotes (Hannoush, 2015). N-Myristoylation commonly occurs cotranslationally, although posttranslational myristoylation was observed during programmed cell death and occurs due to proteolytic cleavage revealing an N-terminal glycine within a cryptic myristoylation consensus sequence, which can then undergo myristoylation (Martin et al., 2011). In contrast, S-palmitoylation is the dynamic reversible addition of a 16-carbon fatty acid, palmitic acid, to cysteine amino acid side chains via a thioester linkage. Targets of this PTM include ion channels, regulatory enzymes, scaffolding proteins, and membrane receptors (Chamberlain and Shipston, 2015). For example, S-palmitoylation of the pro-apoptotic protein BAX regulates its subsequent targeting to the mitochondrial outer membrane to initiate programmed cell death, and the S-palmitoylation of death receptor Fas regulates its protein expression by circumventing its degradation through the lysosome (Fro¨hlich et al., 2014; Rossin et al., 2015). S-Prenylation is another class of lipidation that affects about 2% of the proteome in mammals (Resh, 2006). It is characterized by the irreversible addition of an isoprenoid, either at 15-carbon farnesyl or a 20-carbon geranylgeranyl to one or two C-terminal cysteines of a protein through a thioether linkage. Substrate proteins of S-prenylation require the modification for membrane association and subsequent regulation of function (Berndt et al., 2011; Zhang and Casey, 1996). Much like the other PTMs discussed above, the development of homogeneous synthetic lipopeptides and lipidated proteins have contributed to the biochemical understanding of these modifications. In addition, the creation of a range of chemical probes has transformed the ability to track and identify lipidated proteins from living systems. Encoding Lipidation In cells, short amino acid sequences encode the recognition elements for lipidation and are sufficient enough to promote the modification. In fact, a 10-amino acid sequence transplanted from H-Ras, a known target of S-palmitoylation, to another soluble protein can promote its modification (Hang and Linder, 2011). However, the purification of these proteins in sufficient amounts

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Figure 6. Lipidation (A) Proteins can be modified by several types of lipids, including myristoylation at the N-termini and palmitoylation and prenylation at cysteine residues. (B) Examples of lipid metabolic chemical reporters for the visualization and identification of lipidated proteins and lipid-dependent protein-protein interactions. (C) Acyl-biotin exchange for the analysis of palmitoylation. Free cysteine residues are first capped by incubation of cell lysates with N-ethylmaleimide. Any palmitate thioesters are then cleaved using hydroxylamine and the resulting free thiols can be reacted with a variety of electrophilic tags.

and to homogeneity is still a very difficult task. Synthetic lipopeptides and proteins have given access to fully functional lipidated proteins and have allowed for the study of complete functional proteins. Often the rate-limiting step in these studies is the preparation of lipidated peptides that can be incorporated into larger proteins by techniques like NCL, and accordingly, several different approaches have been developed (Brunsveld et al., 2006). Early work in this area was carried out by Waldmann and coworkers who developed elegant protecting group strategies that enabled the solution-phase synthesis of both base-sensitive palmitate and acid-sensitive farensyl groups (Na¨gele et al., 1998; Schmittberger and Waldmann, 1999; Sto¨ber et al., 1997). SPPS has clear advantages over solution preparation, but it could not be immediately applied to lipidated peptides due to the harsh deprotection and cleavage conditions. In an early effort to circumvent this issue, phenylselenocysteine was introduced into peptides using SPPS and subsequently eliminated under oxidative conditions to give site-specific dehydroalanine residues (Zhu and van der Donk, 2001). These residues can then undergo Michael additions with various thiol-containing nucleophiles, including protected farensyl-thiol. Although this threestep method is attractive due to its ease and versatility, the lack of diastereoselectivity in the reaction is a limitation. Alternatively, on-resin lipidation was used where selectively protected cysteine residues were revealed and alkylated or esterified with farnesyl or palmitoyl electrophiles, respectively (Ludolph et al., 2002). This approach has also been reversed: b-bromoalanine residues were incorporated into a peptide and then displaced by an appropriate thiol-containing lipid as the nucleophile (Pachamuthu et al., 2005). Unfortunately, these methods require a significant excess of the lipids. Notably, the problems with SPPS with pre-lipidated amino acids was first overcome through the use of a hydrazide linker to the solid support, enabling a more straightforward cassette approach (Kragol et al., 2004; Lumbierres et al., 2005). More recently, the concept of post-pep-

tide-synthesis modification has been revised using thiol-ene chemistry (Triola et al., 2008) by selectively reacting cysteine thiols in unprotected peptides with lipid alkenes for the generation of palmitoylation analogs (Calce et al., 2014; Wright et al., 2013) and native farnsylated structures (Calce et al., 2014). Much like the synthesis of other modifications, the generation of analogs of the lipid linkage has also been explored. For example, the Davis laboratory created disulfide-linked lipid modifications by taking advantage of Lawesson’s reagent for the site-selective lipidation of cysteine residues in full-length proteins (Gamblin et al., 2008). Specifically, the cysteine sulfhydryl on the protein is first activated as a phenyl selenenyl sulfide by treatment with phenylselenenyl bromide, followed by addition of lipid thiols, resulting in reasonable modification yields of a model protein (geranyl >90%, farensyl >50%). These peptide synthesis strategies have enabled the generation of lipidated analogs of the protein NRas that were then microinjected into living cells to directly monitor palmitate turnover kinetics and interestingly, enabling the localization of depalmitoylation events throughout the cell, the palmitoylation machinery to the Golgi (Rocks et al., 2010). Decoding Lipidation Classically, protein lipidation has been investigated by treatment of cells with radioactive lipids (3H or 14C) that are metabolically incorporated by cells. However, visualization of the modified proteins requires days-long exposure times on radioactive film, and this technique offers no opportunity for the enrichment or identification of lipidated proteins. To overcome these challenges, fatty acid MCRs containing azides or alkynes have been developed that take advantage of bioorthogonal chemistry in the same way as the carbohydrate reporters discussed above (Figure 6B) (Hang et al., 2011). Specifically, probes are designed to mimic the hydrocarbon chain length, incorporating the clickable substituent at the omega end, therefore minimizing interference with acyl-CoA recognition and enzymatic catalytic

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Cell Chemical Biology

Review Figure 7. Acetylation (A) The most common form of protein acetylation is the dynamic modification of lysine side chains. (B) Acetylated lysine residues can be incorporated into recombinant proteins using unnatural amino acid mutagenesis. (C) Analogs of lysine acetylation can be installed onto cysteine residues using alkylation chemistry, resulting in stable thiocarbamate structures. (D) Chemical reporters of acetylation, malonylation, and aspirin-dependent acetylation.

efficiency (Hannoush, 2015). Recently, for example, Alk-12 (13tetradecynoic acid) was used in conjunction with a biotin-enrichment tag for the identification of myristoylated proteins in the malaria pathogen Plasmodium falciparum, which allowed for the identification of several proteins implicated in the parasite’s life cycle and disease transmission (Wright et al., 2014). Similarly, Alk-16 (17-octadecyonic acid) has been utilized in the proteomic identification and subsequent characterization of palmitoylated proteins. For example, global proteomic profiling in a mouse dendritic cell line identified 150 potentially palmitoylated proteins, including the innate immune effector IFITM3, which the authors subsequently demonstrated requires fatty acylation for its antiviral activity (Yount et al., 2010). Another proteomics study utilized Alk-16 as well as Alk-12 and Alk-14 (i.e., Alk-12 for myristoylation and Alk-16 for palmitoylation) to highlight the selectivity of the chain length for different modifications (Wilson et al., 2011). The chemical reporter Alk-16 has also been used in conjunction with stable isotope labeling in cell culture (SILAC) to perform quantitative proteomics and subsequently demonstrated that some palmitoylation events are stable over time while others are more dynamic (Martin et al., 2012). More recently, a diazirine photo-cross-linking functionality was incorporated into Alk-16, which enabled the identification of proteinbinding partners of lipidated IFITM3 (Figure 6B) (Peng and Hang, 2015). An alternative strategy takes advantage of the thioester linkage of S-palmitoylation to use chemoselective reactions to perform a biotin switch, a method termed acyl-biotin exchange (Figure 6C) (Drisdel and Green, 2004). Specifically, free cysteines in a cell lysate are alkylated with N-ethylmaleimide, followed by treatment with hydroxylamine, which acts to cleave any palmitate thioesters revealing sulfhydryl groups that are then selectively modified with biotinylation reagents. Importantly, this method was used for the identification of palmitoylated proteins in yeast (Roth et al., 2006), mammalian cells (Ivaldi et al., 2012), malaria parasites (Jones et al., 2012), and mouse tissue (Wan et al., 2013). S-Prenylation has also been investigated using alkyne derivatives of isoprenoids (Charron et al., 2011; DeGraw et al., 2010). For example, alkyne-farnesol was utilized in a large-scale enrichment of isoprenoid-modified proteins in a macrophage cell line that led to the identification of both known and unpredicted S-prenylated proteins, including the zinc-finger antiviral protein (ZAP) (Charron et al., 2013).

Other probes developed to investigate palmitoylation machinery take advantage of some irreversible pan palmitoylation inhibitors. For example, alkyne analogs of the inhibitor 2-bromopalmitate (2-BP) were utilized as activity-based probes for protein palmitoyl acyltransferases (PATs) as well as other palmitoylating and 2-BP-binding enzymes (Zheng et al., 2013). Specifically, the corresponding terminal alkyne analogs of 1,2-bromohexadec-15yonic acid (16C-BYA) and 2-bromohexadec-17-ynoic acid (18C-BYA) were synthesized and subsequently used to identify endogenous proteins in HEK293A and the pancreatic cancer cell line PANC1. While 18C-BYA was able to identify three endogenous PATs from HEK293A cells, other acyltransferases and acyl-CoA enzymes were also enriched, as well as many known palmitoylated substrates, raising the possibility that 2-BP could be incorporated into the cellular lipid pool and used as an acyl donor during palmitoylation. Improving on this study, the same authors developed a clickable analog of the natural product cerulenin, an inhibitor of fatty acid biosynthesis and protein palmitoylation that acts through irreversible alkylation of the cysteine residues in the enzymes (Zheng et al., 2015). The cerulenin-derived probe was demonstrated to be more specific than the first generation 2-BP probe, perhaps because it does not require metabolic transformation within the cell. The Tate laboratory utilized a potent and specific human N-myristoyltransferase (NMT) inhibitor in combination with SILAC and an alkyne-containing myristate analog to identify potentially myristoylated proteins (Thinon et al., 2014). In brief, cells were treated with tetradec-13-ynoic acid (YnMyr) (Heal et al., 2008) and were either grown in standard medium containing NMT inhibitor or in SILAC medium with no inhibition. Following enrichment using an azido-biotin affinity tag, substrates (and non-substrates) were assigned according to the response in enrichment to the inhibition of NMT. Using this method, over 100 N-myristoylated proteins were identified including novel targets such as nucleolar protein 3 (NOL3/ARC), a protein implicated in inhibition of apoptosis, tumorigenesis, metastasis, and chemoresistance (Thinon et al., 2014). Acetylation Protein acetylation describes the reversible posttranslational transfer of an acetyl group from acetyl-coenzymeA (acetylCoA) to a target protein, most commonly on the side-chain amine of a lysine residue (Figure 7A). This process is catalyzed by lysine acetyltransferases (KATs) and is removed by lysine deacetylases (KDACs) (Yang et al., 2011). KDACs can be broken

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Review into two families: classic metallohydrolases that act directly to hydrolyze acetimide and the sirtuins that utilize an nicotinamide adenine dinucleotide (NAD) cofactor (Dancy et al., 2012). Protein acetylation was first identified on lysine-rich N-terminal tails of histones isolated from calf thymus in 1963 (Phillips, 1963). Approximately 1 year later, Vincent Allfrey and colleagues showed that radiolabeled acetate was rapidly sequestered from medium and incorporated onto histones of isolated nuclei while not being affected by treatment with puromycin, a translation inhibitor, suggesting that the acetylation events take place posttranslationally. In addition, Allfrey was able to show that histone acetylation decreased inhibition of RNA synthesis, giving rise to the widely accepted theory that posttranslational histone acetylation serves as a dynamic and reversible mechanism for the regulation of transcription (Allfrey et al., 1964). Since Allfrey made his initial discovery, his theory has been thoroughly validated, and it is currently accepted that lysine acetylation, in particular that of histones, plays a large role in regulating epigenetic changes through both gene transcription (Shahbazian and Grunstein, 2007) and non-chromatin associated proteins (Yang et al., 2010). Given that changes in transcriptional regulation are a feature of human diseases, most notably cancer, there has been significant interest in the modulation of protein acetylation levels as a therapeutic strategy. For example, vorinostat, a histone deacetylase (HDAC) inhibitor, is the first US Food and Drug Administration-approved drug for acetylation regulation for the treatment of T-cell lymphoma, and vorinostat and two other HDAC inhibitors, romidepsin and panabinostat, are currently going through clinical trials for the treatment of other cancers, as well as HIV infection (Shirakawa et al., 2013; Verdin and Ott, 2015). Despite these successes and the creation of several anti-acetylation antibodies, deciphering the effects of the thousands of known human acetylation sites remains an obstacle, and a variety of chemical tools have been developed to help move the field forward. Encoding Acetylation Examination of site-specific acetylation events is needed to fully understand the biological implications associated with the modification. In contrast to the modifications discussed above, the incorporation of acetylated lysine residues into peptides using SPPS is quite straightforward. Lysine residues directly bearing an ε-N-acetate group can be used without the need for further protection, and then these peptides can readily participate in NCL reactions for the preparation of synthetic proteins (He et al., 2003). For example, NCL was used to generate histone H4 site specifically acetylated at K16 (Shogren-Knaak et al., 2006). Subsequent biochemical analysis showed that acetylation played an integral role in chromatin compaction through inhibition of cross-fiber interactions. They also found that this acetylation event inhibits ATP-dependent chromatin assembly and remodeling enzyme (ACF) from mobilizing the mononucleosome, suggesting that acetylation of K16 is sufficient to regulate both higher-order chromatin structure as well as protein-chromatin interactions in a manner that affects its function (Shogren-Knaak et al., 2006). In addition to NCL-based approaches, acetyl-lysine has also been incorporated into recombinant proteins using unnatural amino acid mutagenesis (Figure 7B) (Neumann et al., 2008, 2009). The Megachile bakeri pyrrolysyl-tRNA synthetase/tRNACUA was once again used to incorporate acety-

lated lysine at amber stop codons in E. coli expressed proteins for biochemical experiments. For example, acetyl-lysine was site specifically introduced at residue 56 in histone H3, and single-molecule fluorescence resonance energy transfer experiments were used to show that it does not have a direct effect on the compaction of chromatin (Neumann et al., 2009). Finally, an analog of acetyl-lysine has also been encoded through modification of cysteine residues (Figure 7C) (Huang et al., 2010). Specifically, Cole and coworkers demonstrate that cysteine residues in peptides and proteins can be selectively alkylated in high yield with methylthiocarbonyl-aziridine to generate a thiocarbamate analog of acetyl-lysine. Importantly, this analog is recognized by both antibodies and an acetyl-lysine-binding bromodomain, and the authors confirmed its ability to mimic the natural modification in the activation of two full-length proteins. Notably, this analog is stable to enzymatic deacetylation, potentially enabling the specific effects of an acetylation mark to be tested in cell lysates or through microinjection. Decoding Acetylation Although anti-acetyl-lysine antibodies exist, chemical reporters of acetylation have certain advantages, including reporting on non-lysine acetylation events and robust recovery in proteomics experiments. Toward this goal, sodium 4-pentynoate has been developed as an MCR lysine acetylation both in living cells and in vitro (Figure 7D). Treatment of living cells with 4-pentynoate or cell lysates with chemically synthesized 4-pentynoyl-CoA enabled the visualization and identification of both known and new acetylation targets (Yang et al., 2010). Subsequently, 4-pentynoyl-CoA was incubated with the acetyltransferase p300 to identify enzyme-specific substrates. The newly discovered acylation substrates included a cysteine residue in histone H3 variants (Wilson et al., 2011), demonstrating that these reporters can be used to visualize acylation events that would not be picked up by traditional anti-acetyl-lysine antibodies. Additional acylation events with diverse structures on lysine residues have also begun to emerge. To further probe these understudied modifications, an alkyne-bearing MCR of lysine malonylation was synthesized as a protected version of 2-propargyl malonate, termed Mal-AMyne (Figure 7D) (Bao et al., 2013). Treatment of HeLa cells with Mal-AMyne resulted in the identification of 14 previously known malonylated proteins as well as 361 new potential substrates (Bao et al., 2013). Another form of acylation that occurs is a non-enzymatic transfer of acetate from small molecules, such as aspirin, to protein substrates. Our laboratory introduced a chemical reporter of aspirin acetylation, AspAlk (Figure 7D) that enabled the visualization of these chemical events and identification of 120 potential substrate proteins including some of the core histones (Bateman et al., 2013). More recently, this same probe was combined with quantitative proteomics to identify 523 proteins and many specific sites of aspirin-mediated acetylation (Wang et al., 2015). Methylation Protein methylation describes the transfer of a methyl group onto the side chains of lysines, arginines, and less commonly histidines (Figure 8A) (Biggar and Li, 2015; Lee et al., 2005). Lysine residues can be mono-, di-, or tri-methylated, while arginines can be mono-methylated or di-methylated. Histidines have only been reported to be mono-methylated, however, this

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Review Figure 8. Methylation and ADP-Ribosylation (A) A variety of methylation marks can be dynamically installed onto both lysine and arginine side chains. (B) Mono-methylation can be incorporated into recombinant proteins using unnatural amino acid mutagenesis followed by acid-based deprotection. (C) Mono-, di-, and tri-methylated lysine analogs can be generated by alkylation of cysteine residues with the appropriate ethyl-amino electrophile. (D) Using a bump-hole approach, the alkynebearing SAM analog will be transferred by engineered methyltransferases, enabling the identification of transferase-specific substrates. (E) ADP-ribose can be enzymatically added to a variety of protein side chains and then subsequently polymerized to form long poly-ADP-ribose chains. (F) ADP-ribose analogs can be installed onto peptides after solid-phase peptide synthesis by taking advantage of oxime chemistry. (G) Examples of ADP-ribosylation reporters for use in cell lysates.

modification is uncommon and its role remains unclear (Greer and Shi, 2012). Histone methylation plays well-documented roles in transcriptional regulation, and non-histone protein methylation, although historically less characterized, has emerged as a prevalent PTM that plays an important role in cellular signaling. The installation of posttranslational methyl modifications onto proteins is catalyzed by a class of enzymes called methyltransferases and is removed by demethylases. Methyltransferases generally use S-adenosylmethionine (SAM) as a methyl donor (Biggar and Li, 2015). Demethylases use a FAD cofactor and molecular O2 to produce formaldehyde, hydrogen peroxide, and the demethylated peptide (Dancy et al., 2012). Protein methylation on lysine was first recognized to occur posttranslationally in 1965 (Kim and Paik, 1965), shortly after methylated lysine was found in bacterial flagellar protein

(Ambler and Rees, 1959) and on the histones isolated from calf thymus, wheat germ, and multiple rat organs (Murray, 1964). Since then, protein methylation has been implicated in a variety of cellular functions including cell-cycle regulation, DNA damage and stress response, and the development and differentiation through modulation of chromatin-bound histones as well as chromatin-associating non-histone proteins (Huang and Berger, 2008; Kouzarides, 2007). Not surprisingly, the misregulation of protein methylation has been linked to a variety of diseases including cancer (Chi et al., 2010), intellectual disability (Iwase and Shi, 2010), and aging (Scaffidi and Misteli, 2006). Antibodies that recognize the different methylation states of both lysine and arginine are available, as well as small-molecule inhibitors of methyltransferases and demethylases. These tools have contributed greatly to the investigation of protein methylation, but these methods have limitations for the analysis of specific methylation events. Fortunately, chemical approaches for the preparation of site specifically modified proteins, as well as probes that can deconvolute the substrate specificity of methyltransferases, have both contributed to our understanding of this key modification. Encoding Methylation Like acetylation, methylated lysine residues can be easily incorporated into peptides using SPPS; di-methyl and tri-methyl lysines can be directly incorporated, and the mono-methylated side chain can be protected as an appropriate carbamate. NCL has been frequently used for incorporating methylated lysine with native linkages into proteins (He et al., 2003). In an excellent example of the power of NCL, a mononucleosomes bearing either a di- or tri-methylated lysine (K7) histone H3 and

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Review a series of acetylated lysine residues in histone H4 were prepared (Ruthenburg et al., 2011). Using these homogeneous proteins, the authors were able to demonstrate the cross-talk that occurs between the two PTMs by showing a significant increase in binding by the BPTF PHD-bromodomain in response to a specific pattern of both types of modification. In another example, NCL was used to generate histone H3 with tri-methylated lysines at residues 4, 9, and 27 (Bartke et al., 2010). This semisynthetic methylated H3 was combined with biotinylated DNA and immobilized on streptavidin beads to enrich for binding partners of H3 that are specific for the tri-methylation modifications. As a complementary strategy, unnatural amino acid mutagenesis using the M. bakeri pyrrolysyl-tRNA synthetase/tRNACUA has also been used to incorporate mono-methylated lysine residues (Figure 8B). A pyrrolysyl-tRNA synthetase pair has been used extensively for incorporation of mono- and di-methylated lysine into recombinant proteins as it prefers methylated lysine over unmodified lysine. In one example, mono- and di-methylated Lys were incorporated into recombinant proteins using a PylRstRNA pair. Here, mono-methylated lysine was installed as a tert-butoxycarbonyl protected analog, which was then subsequently deprotected using TFA (Nguyen et al., 2009). Similar two-step genetic approaches have been used for the installation of site-specifically modified lysine using alternative protection groups including allylcarbamoyl and photocaged methylated lysine that allow for alternative deprotection methods (Ai et al., 2010; Groff et al., 2010; Wang et al., 2010). The Chin laboratory has also used a GOPAL-like strategy to reveal a specific lysine residue that can be di-methylated using reductive methylation (Nguyen et al., 2010). Unnatural amino acid mutagenesis methods are yet to be used for the direct incorporation of diand tri-methylated lysine residues. However, a chemical method for the facile incorporation of mono-, di, and tri-methylated lysine analogs was developed (Figure 8C) (Simon et al., 2007). Briefly, cysteine resides at the desired site of modification can be selectively alkylated to create a methyl lysine analog (MLA), N-methylated aminoethylcysteine. Notably, these mono-, di-, and tri-methylated lysine analogs were functionally similar to their native counterparts when incorporated into recombinant proteins. More recently, a similar strategy was applied to arginine methylation through the reaction of cysteine residues with a,bunsaturated amidines to generate both mono- and di-methylated arginine analogs (Le et al., 2013). Decoding Methylation Linking a specific methyltransferase to a particular substrate is key to understanding the complex cellular biology of this modification. Working to identify and analyze the protein methylome, two alkyne analogs of SAM were developed as the first chemical reporters of protein methylation. Several endogenous methyltransferases accepted these stable alkyne analogs of SAM as cofactors and were able to transfer an alkylated methyl group onto a lysine residue of both a peptide and a recombinant protein. The alkylated methyl group could then subsequently be subjected to labeling with CuAAC using an azide tag (Binda et al., 2011; Peters et al., 2010). Interestingly, the alkyne-SAM analogs were utilized selectively by different methyltransferases, suggesting that a chemical reporter/methyltransferase pair could be created in the same way that selective reporters of phosphor-

ylation were developed as described above. Toward this goal, an additional series of azide- and alkyne-SAM analogs of various sizes were prepared (Luo, 2012). Several of the SAM analogs developed were not turned over by wild-type methyltransferases but were selective for rationally engineered methyltransferase mutants using the bump-hole strategy, enabling the identification of the specific methyltransferase-dependent substrates (Figure 8D) (Islam et al., 2013). Although this strategy was useful for in vitro testing and the screening of cellular extracts, the poor cell permeability of SAM analogs limited their use in experiments involving living cells. To overcome this, the biosynthetic pathway for SAM was engineered in mammalian cells (Wang et al., 2013b). Briefly, cells can be treated with cell-permeable alkyne-methionine analogs that will be enzymatically transformed to the corresponding SAM analogs. Due to the physiological instability of these methionine-based SAM analogs, a more stable seleniumbased SAM reporter was also created that can probe both arginine and methyltransferases in vitro (Willnow et al., 2012). Continuing the work with selenium-based reporters, it was shown that a propargylic Se-containing SAM analog could be used by endogenous methyltransferases and was stable in whole-cell lysates (Bothwell et al., 2012). ADP-Ribosylation Protein ADP-ribosylation describes the posttranslational transfer of an ADP-ribose moiety from b-nicotinamide adenine dinucleotide (NAD+) to a variety of amino acid side chains on protein acceptors, including aspartate, glutamate, lysine, arginine, and cysteine (Figure 8E) (Daniels et al., 2015; Leung, 2014). This mono-ADP-ribosylation is then often polymerized to generate a long chain of repeating units, termed poly-ADP-ribosylation. In humans, ADP-ribosylation is installed by a family of 17 diphtheria toxin-like ADP-ribosyltransferases (ARTDs), commonly known as poly-ADP-ribose polymerases (PARPs). The majority of these enzymes catalyze mono-ADP-ribosylation, and four PARPs (PARP1, 2, 5a/b) are known to catalyze poly-ADP-ribosylation through the transfer of multiple ADPr units onto target proteins (Carter-O’Connell et al., 2014; Morgan and Cohen, 2015). ADP-ribosylation can be removed by endogenous enzymes that cleave poly-ADP-ribose polymers such as poly(ADP-)ribose glycohydrolase (Moyle and Muir, 2010). With NAD+ serving as a substrate for ADP-ribosylation, NAD+ consumption and energy metabolism are directly linked to the production of ADP-ribose derivatives (Schreiber et al., 2006). Like many PTMs, ADP-ribosylation plays a unique role in many important cellular processes including but not limited to stress signaling, DNA damage repair, telomere homeostasis, transcriptional regulation, and centrosomal targeting (Carter-O’Connell et al., 2014; Schreiber et al., 2006). ADP-ribosylation has also been shown to have important therapeutic consequences in cancers, neurodegenerative diseases, ischemia, and inflammatory disorders (Curtin and Szabo, 2013). The specific function of the majority of ribosylation events, in particular mono-ADP-ribosylation, is not well understood. Obstacles contributing to difficulties in studying ADP-ribosylation include the stability of the ester-linked ADP-ribose at basic pH, the ability of the modification to be rapidly removed by endogenous enzymes, lack of commercial antibodies, and overlapping target specificities among the 17 ARTDs (Carter-O’Connell et al., 2014; Moyle and Muir, 2010). Again, chemical methods

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Review have begun to address these limitations through the site-specific installation of ADP-ribose analogs and the development of chemical probes for the global and isoform-specific identification of ARTD substrates. Encoding ADP-Ribosylation As with the more extensively studied modifications above, the preparation of site-specifically ADP-ribosylated peptides and proteins is key to investigating the biochemical effects. However, the chemically sensitive nature of the pyrophosphate bond makes peptide synthesis challenging. In order to circumvent these issues, Filippov and coworkers installed a selectively protected ribosylated asparagine or glutamine residue into peptide using SPPS (van der Heden van Noort et al., 2010). An alternative approach has also been developed for the generation of ADP-ribosylation analogs (Figure 8F) (Moyle and Muir, 2010). More specifically, this method uses aminooxy-functionalized amino acids for the specific conjugation of ADP-ribose onto peptides and semisynthetic proteins, with oxime ligation occurring between the aminooxy-functionalized amino acid of choice and the anomeric carbon of ribose at pH 4.5. Notably, this mimic of mono-ADP-ribosylation in an H2B-derived peptide was a substrate for PARP1, producing site-specific-poly-ADP-ribosylated peptide conjugates. Finally, the same authors used NCL to prepare ADP-ribosylated H2B proteins with benzophenone cross-linkers and were able to enrich for previously unknown ADP-binding proteins, histone mH2A1.1 and PARP9 (Moyle and Muir, 2010). Decoding ADP-Ribosylation In an effort to enrich and identify the ADP-ribosylated proteome, the alkyne-containing chemical reporters 6-alkyne-NAD and 8-alkyne-NAD have been synthesized (Du et al., 2009; Jiang et al., 2010). Incubation of these analogs with cell lysate and recombinant PARP1 led to the discovery of 70 potentially new PARP1 protein substrates (Figure 8G) (Jiang et al., 2010). Orthogonal NAD variants have also been used in combination with several engineered PARPs to identify direct substrates of specific members of the PARP superfamily (Carter-O’Connell et al., 2014). A bump-hole strategy was used to create a mutant ARTD (K903A) that could accept an NAD analog containing an ethyl substituent at the C50 position of the nicotinamide moiety of NAD. Incubation of these orthogonal pairs in nuclear extracts resulted in the proteomic identification of a pool of substrates specific to either PARP1 or PARP2. Affinity purification and MS/MS allowed for identification of unique targets for both enzymes, which could be further applied to all 17 members of the PARP superfamily to delineate the role of specific enzyme-substrate interactions. More recently, an aminooxy alkyne (AO-alkyne) probe was synthesized to detect monoADP-ribosylation in cells using CuAAC and an azide-containing tag. The probe was used to show that PARP10 and PARP11 are auto-ADP-ribosylated as well as used to monitor stimulus-induced ADP-ribosylation in cells (Morgan and Cohen, 2015). Conclusions and Future Outlook Chemical techniques have been particularly valuable for the synthesis and identification of posttranslationally modified proteins. Advances in protein-peptide ligations for the semisynthesis of modified target proteins has enabled the development of unique

and site-specific chemical reactions for the installment of modifications through either native or non-native linkages, further expanding our toolbox. For example, in a tour-de-force of NCL mediated synthesis, the Muir laboratory has recently prepared histones with different combinations of ubiquitination, acetylation, and methylation marks (Nguyen et al., 2014). These proteins can be combinatorially combined to form mononucleosomes with different modification patterns resulting in a PTM library, which, paired with DNA sequencing techniques, can rapidly identify specific protein-binding partners for different patterns of modifications. In parallel, improvements in the quality of chemical reporters and MS methods have allowed for an unprecedented volume of identified proteins and, coupled with chemical tools developed for the enrichment of modified proteins, has resulted in new protein targets for research and drug discovery. Specifically, the application of chemical reporters for the incorporation of bioorthogonal chemical moieties has enabled the study of a variety of posttranslational modifications that, due to their chemical structure and complex regulation, have been difficult to study with traditional biological methods. For example, Hang and coworkers used the Alk-16 MCR to show that changes in the palmitoylation of specific proteins control the entry into meiosis in fission yeast, suggesting that single enzymes that install PTMs can control complex biological processes (Zhang et al., 2013). Unfortunately, despite these successes, challenges still remain. For example, in some cases, the use of SPPS for the synthesis of modified peptides relies on the chemical synthesis of complex pre-modified amino acids that require unique protecting group strategies, limiting its use to laboratories with the requisite chemical expertise. Furthermore, NCL suffers from concentration-dependent reaction rates, which limits its utility with folded protein substrates that cannot be concentrated to reasonable levels. Therefore, there is a need for continued collaboration between chemists and biologists to devise recombinant strategies for the preparation of evermore challenging protein targets. In the case of chemical reporters, treatment with metabolic analogs may perturb the cellular environment, causing unnatural changes in cellular metabolism and altering metabolic pathways. This highlights the need for a more comprehensive investigation, including understanding the cellular fate of these analogs to uncover which PTMs are being enriched with a specific reagent. Beyond the identification of modified substrate proteins, retrieving site-specific information still remains a challenge, as many PTMs are not stable during MS/MS analysis. These shortcomings should be met with a research effort that focuses on enhanced MS methods, including new ionization techniques and computer programs that allow for the direct identification of modification sites. Chemical methods have been instrumental in the investigation of PTMs toward elucidating their biological role, and we believe that certain modifications, such as arginine methylation, are particularly ripe for the development of additional chemical tools. The exciting advancements in this field have enabled the use of multiple chemical methods to both synthesize homogeneous proteins for study while unambiguously identifying their modification status, and undoubtedly, chemical biologists will continue to have a huge impact in the field of protein modifications.

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Review AUTHOR CONTRIBUTIONS

Berndt, N., Hamilton, A.D., and Sebti, S.M. (2011). Targeting protein prenylation for cancer therapy. Nat. Rev. Cancer 11, 775–791.

K.N.C., A.R.B., and M.R.P. wrote the manuscript and prepared the figures.

Biggar, K.K., and Li, S.S.C. (2015). Non-histone protein methylation as a regulator of cellular signalling and function. Nat. Rev. Mol. Cell Biol. 16, 5–17.

ACKNOWLEDGMENTS

Binda, O., Boyce, M., Rush, J.S., Palaniappan, K.K., Bertozzi, C.R., and Gozani, O. (2011). A chemical method for labeling lysine methyltransferase substrates. Chembiochem 12, 330–334.

K.N.C. is a fellow of the National Science Foundation Graduate Research Fellowship Program (DGE-0937362). Our current research is supported by the National Institute of General Medical Sciences (R01GM114537), the National Science Foundation (CHE-1506503), Susan G. Komen for the Cure (CCR14299333), and the American Cancer Society (RSG-14-225-01-CCG). The authors thank Cesar De Leon for help in the preparation of Figure 1. REFERENCES

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