Chemical Methods for Encoding and Decoding of Posttranslational Modifications.

HHS Public Access Author manuscript Author Manuscript

Cell Chem Biol. Author manuscript; available in PMC 2017 January 21. Published in final edited form as: Cell Chem Biol. 2016 January 21; 23(1): 86–107. doi:10.1016/j.chembiol.2015.11.006.

Chemical methods for encoding and decoding of posttranslational modifications Kelly N. Chuh1, Anna R. Batt1, and Matthew R. Pratt1,2,3 1Department

of Chemistry, University of Southern California, Los Angeles, CA 90089

2Department

Author Manuscript

of Molecular and Computational Biology, University of Southern California, Los Angeles, CA 90089

Abstract

Author Manuscript

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 compliment 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 chemistrybased 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

Author Manuscript

In the post-genomic 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

3

Correspondence should be addressed to [email protected]. Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. Author Contributions K.N.C., A.R.B., and M.R.P. wrote the manuscript and prepared the figures.

Chuh et al.

Page 2

Author Manuscript Author Manuscript

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 often requires tedious and often difficult separation of their modified and unmodified forms. Furthermore, PTMs can occur on several sites simultaneously and substoichiometricly, making the isolation of a completely homogenous population extremely difficult. Therefore, access to site-specifically modified proteins is of the utmost importance for the study of PTMs. Additionally, 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.

Author Manuscript

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 in PTMs that have only been addressed by one approach, we direct readers to other excellent reviews (Chuh and Pratt, 2015a; L. Davis and Chin, 2012; Grammel and Hang, 2013; Muir, 2003; Vila-Perelló and Muir, 2010).

Phosphorylation

Author Manuscript

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 1A). The modification is installed by members of the kinase family of enzymes, which transfer the high-energy gamma phosphate from adenosine triphosphate (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 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; J. Zhang et al., 2009), neurodegeneration (L. H. Wang et al., 2004), and diabetes (Prada and Saad, 2013; Winder and Hardie, 1999). Accordingly, there is

Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 3

Author Manuscript

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 (CML) (Capdeville et al., 2002). This has been followed by the clinical development of other inhibitors of growth factor receptor kinases, Bruton’s tyrosine kinase (BTK) (Pan et al., 2007), and others. Despite these exciting results, gaining a complete understanding of the many roles of phosphorylation remains extremely challenging.

Author Manuscript

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 1B) (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 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 acyl-transfer, 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 1C) (Vila-Perelló and Muir, 2010). Specifically, proteins 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 to commercially available intein constructs (N. H. Shah et al., 2012; N. H. Shah and Muir, 2013).


Chuh et al.

Page 4

Author Manuscript Author Manuscript Author Manuscript Author Manuscript

Phosphorylated tyrosine, serine, and threonine residues and their corresponding nonhydrolyzable 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 Cterminus 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 a intramolecular interaction between the phosphorylated tail and SH2 domain in Src. Using EPL, the authors prepared a 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 towards 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 TGFβ signaling pathway (Massagué, 2012). Extracellular TGFβ will engage with TGFβ receptor kinase (TGFβR), 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 TGFβR 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; Z. 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 (CK2α) with a difluoro-phosphonate at threonine residue 344. A subsequent combination of microinjection and in vitro assays demonstrated that this modification increases the cellular stability of CK2α and alters its substrate selectivity (Tarrant et al., 2012). More recently, Lashuel and coworkers used EPL to prepare both unmodified and phosphorylated versions the Huntingtin exon 1 protein that is prone to aggregation that causes Huntington’s disease (Ansaloni et al., 2014). Using these


Chuh et al.

Page 5

Author Manuscript

proteins, the author showed that phosphorylation at threonine residue 3 significantly slows the aggregation of this protein.

Author Manuscript Author Manuscript Author Manuscript

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, which 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 1D). 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 a orthogonal codon that can be read out by the ribosome during protein translation (Chin, 2014; Lang and Chin, 2014; C. C. Liu and Schultz, 2010). Dozens of unnatural amino acids have now been incorporated into proteins in E. 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 two-step 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 could be engineered to favor incorporation of phosphorylated serine into recombinant proteins. This was done and worked as expected (S. 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 EF-Tu. 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. Additionally, 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


Chuh et al.

Page 6

Author Manuscript

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 has enabled the global characterization of the phosphoproteome. 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 2A) (Y. Liu et al., 1998; K. Shah et al., 1997; Ubersax et al., 2003; C. 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 gate-keeper residue prevents the binding of an N6-benzyl-modified ATP in the kinase active site. However, when the gate-keeper 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 ATP-competitive 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 ATPγS analog to thiophosphorylate proteins was exploited (Allen et al., 2007; 2005). In this elegant strategy, the resulting protein thiophosphate can then be alkylated with a p-nitrobenzyl group (Figure 2B). An antibody that specifically recognizes this alkylated structure can then be used to either visualize or enrich kinase substrates for proteomic identification. Combining analog-sensitive kinases with this approach allows for the enrichment and identification of the substrates of in principal 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 threecomponent reaction between a substrate protein (or peptide) of interest, the small-molecule,


Chuh et al.

Page 7


and the conserved active-site lysine residue of a kinase (Maly et al., 2004; Riel-Mehan 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 2C). 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 photocross-linking groups into an analog of ATP (Parang et al., 2002). Upon photo-irradiation, one of the cross-linkers near the adenosine ring will covalently label 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 endoplasmic reticulum (ER) and Golgi, resulting in Nlinked 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 α-O-linkage to the β-hydroxyl group of serine or threonine residues. This monosaccharide can then be elaborated through addition of sialic acid, fucose and/or additional units of Galβ1,4GlcNAc 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 dolichollinked 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-acetylglucosamine (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


Chuh et al.

Page 8

Author Manuscript

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. Further more, 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 (B. 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 α-O-linked “cassettes” for solid phase peptide synthesis (Figure 3A). For example, the Boons lab 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 α-OSer/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 lab 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 3B). For example, the Keissling lab 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 (Y. He et al., 2004). More recently, Doores and coworkers have demonstrated a 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 N-linked glycosylation during protein folding in the ER, Kajihara and Ito synthesized misfolded, interleukin-8 (IL-8, CXCL8) bearing an N-linked glycan. Using this material, they found that the enzyme UDPglucose:glycoprotein glucosyltransferase (UGGT), prefers misfolded over correctly folded glycoproteins, indicating that UGGT plays a distinct role in quality control of protein


Chuh et al.

Page 9

Author Manuscript

folding (Izumi et al., 2012). 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 (P. Wang et al., 2013). Our lab has utilized EPL to study the function of O-GlcNAc modification on the protein α-synuclein, the major aggregating protein associated with Parkinson’s disease and dementia with Lewy Bodies (DLBs). Chemical synthesis of homogenous, full-length, O-GlcNAcylated αsynuclein was achieved using EPL in three steps and subsequent aggregation assays showed that O-GlcNAc modification at threonine residue 72 inhibits α-synuclein aggregation and is non-toxic to neurons in cell culture (Marotta et al., 2015).


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. Additionally, some glycosidic bonds cannot survive the strong acidity of final deprotection conditions of peptides (e.g. TFA). To address these shortcomings, chemoenzymatic methods to catalyze the addition of single monosaccharides onto to 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 and co-workers recently demonstrated the stepwise, chemoenzymatic synthesis of O-linked glycosylated MUC1 peptides utilizing Drosophila glycosyltransferases (Bello et al., 2014). In the area of N-linked glycoproteins, the Boons and Paulson labs 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 (Z. Wang et al., 2013). The resulting library was printed on a microarray and was subsequently screened for binding to lectins and influenzavirus 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-β-N-acetylglucosaminidases (ENGases) have been utilized to perform transglycosylation reactions (Figure 3C) (L.-X. Wang, 2011). These enzymes normally cleave N-linked glycans, leaving only the first N-acetylglucosamine (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 the Nlinked 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 α-1,6-fucosylated GlcNAc derivatives, producing native, core fucosylated, complex-type glycopeptides (W. Huang et al., 2011). More recently, the Davis lab has explored a bacterial endoglycosidase, EndoS, that is complimentary to other


Chuh et al.

Page 10

Author Manuscript

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, pre-formed 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 branch points, as oximes (Rodriguez et al., 1997) and thioethers respectively (Marcaurelle and Bertozzi, 2001), as well as thioether linkages to install antennae onto a 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 and co-workers 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 (Dondoni et al., 2009). 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-and-modify” 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 nonnatural 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 (Fernández-González et al., 2010). In a similar fashion, an alkyne-containing nonnatural amino acid, homopropargylglycine (Hpg) can be incorporated as a methionine surrogate. Interestingly, the Davis lab utilized both Aha and Hpg to enable attachment of multiple glycans to bacterially expressed protein scaffolds (van Kasteren et al., 2007). Decoding protein glycosylation

Author Manuscript

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 lab, several labs have investigated the use of metabolic chemical reporters (MCRs) for the visualization and identification of glycosylated proteins (Figure 3D). 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 lab (Saxon and Bertozzi, 2000). Once this compound diffuses into cells, the O-acetates are removed by endogenous lipases and the resulting ManNAz is biosynthetically


Chuh et al.

Page 11


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 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 lab subsequently found that the alkyne MCR, Ac4GlcNAlk, displayed improved signal to noise and was more selective for O-GlcNAc modifications compared to 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 a 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; Späte et al., 2014) and cyclopropenes (Cole et al., 2013; Patterson et al., 2014; 2012), 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 C. elegans (Laughlin and Bertozzi, 2009). Additionally, 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 lab has developed a technique termed isotope-targeted glycoproteomics (IsoTaG), a mass-independent chemical glycoproteomics method for the identification of intact, metabolically labelled (using Ac4ManNAz or Ac4GalNAz) glycopeptides from the total proteome (Woo et al., 2015). In contrast to traditional tandem 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 lab 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 an recombinantly expressed, mutant β-1,4-galactosyltransferase and chemically prepared UDPGalNAz results in the transfer of GalNAz to O-GlcNAc residues (Figure 3E) (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). Additionally, this strategy has been combined


Chuh et al.

Page 12

Author Manuscript

with both β-elimination (Khidekel et al., 2007) and electron transfer dissociation mass spectrometry [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 4A) (Z. J. Chen and Sun, 2009; Komander and Rape, 2012). Ubiquitin is added 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) Akin to phosphorylation, ubiquitylation 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, the 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

Author Manuscript

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 co-workers 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 4B) (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, thiolbearing ligation auxiliary attached to the ε-amino group of Lys120 through a glycine linker


Chuh et al.

Page 13


(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 by irradiation with ultraviolet light. A second EPL reaction was preformed between this ubiquitinated peptide and a recombinant H2B(1–116) C-terminal thioester to give fulllength, 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 δ-mercaptolysine unnatural amino acid building blocks for solid phase peptide synthesis first developed by the Brik lab (Ajish Kumar et al., 2009; Kumar et al., 2010) followed shortly by Ovaa and coworkers (Figure 4C) (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 α-synuclein was prepared bearing either mono-, di-, or tetraubiquitination at the physiologically-relevant lysine residue 12 and the different effects on the protein aggregation and stability was 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 labs 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 mutagenesis 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 lab designed a synthetic scheme in which a δ-thiol-L-lysine is incorporated at the desired site of ubiquitylation using an improved pyrrolysyl-tRNA synthetase/tRNACUA pair (Figure 4D) (Virdee et al., 2011). Using this method, Chin and co-workers prepared K6linked 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 semi-synthesis of ubiquitinated proteins. To address this shortcoming, several groups have created strategies to introduce ubiquitin onto substrate


Chuh et al.

Page 14


proteins through isopeptide bond mimics (Figure 4E). One of the first such methods was a disulfide-directed approach that was developed simultaneously by the Muir and Zhuang labs (Chatterjee et al., 2010; J. Chen et al., 2010). This approach takes advantage of a ubiquitinintein 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 reagents like 2,2′-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 lab took advantage of thiol-ene coupling (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 wild-type isopeptide bond, demonstrating that it is a good structural mimic. In yet another thioether approach, the Brik lab utilized either an α-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 alkyne-containing amino acids (Eger et al., 2011; Rösner et al., 2015; Sommer et al., 2011). Additional methods based on the use of nonnatural linkages continue to be developed and applied, including propanone (Yin et al., 2000) and oxime (Shanmugham et al., 2010) linkages. Decoding protein ubiquitylation

Author Manuscript

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, DUBspecific 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


Chuh et al.

Page 15

Author Manuscript

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 carboxyl-terminal hydrolase isoenzyme L3 (UCHL3). Confirmed by X-ray crystallography, it was found that the resulting quaternary vinyl thioether conferred selectivity towards 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 5A) (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 under go myristoylation (D. D. O. 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 Spalmitoylation of death receptor Fas regulates its protein expression by circumventing its degradation through the lysosome (Fröhlich 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; F. L. 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. Additionally, the creation of a range of chemical probes has transformed the ability to track and identify lipidated proteins from living systems.


Chuh et al.

Page 16

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 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 (Nägele et al., 1998; Schmittberger and Waldmann, 1999; Stöber et al., 1997). Solid-phase peptide synthesis 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 solid-phase peptide synthesis 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 three-step method is attractive due to its ease and versatility, the lack of diastereoselectivity in the reaction is a limitation. Alternatively, onresin 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: β-bromo-alanine 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 solid-phase peptide synthesis with pre-lipidated amino acids was first overcome through the use of a hydrazide linker to the solid support, enabling a more straight-forward cassette approach (Kragol et al., 2004; Lumbierres et al., 2005). More recently, the concept of post-peptide-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; T. H. 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 lab created disulfide-linked lipid modifications by taking advantage of Lawesson’s reagent (LR) 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% while 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). Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 17

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 5B) (Hang et al., 2011). Specifically, probes are designed to mimic the hydrocarbon chain length, incorporating the “click-able” substituent at the omega end, therefore minimizing interference with acyl-CoA recognition and enzymatic catalytic efficiency (Hannoush, 2015). Recently, for example, Alk-12 (13-tetradecynoic 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 (M. H. 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 anti-viral 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 (B. R. Martin et al., 2012). More recently, a diazirine photo-cross-linking functionality was incorportated into Alk-16, which enabled the identification of protein-binding partners of lipidated IFITM3 (Figure 5B) (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 (ABE) (Figure 5C) (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 Sprenylated 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 2bromopalmitate (2-BP) were utilized as activity-based probes for protein palmitoyl acyltransferases (PATs) as well as other palmitoylating and 2-BP-binding enzymes (Zheng


Chuh et al.

Page 18


et al., 2013). Specifically, the corresponding terminal alkyne analogs of 1,2bromohexadec-15-yonic 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 2BP 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 lab utilized a potent and specific human Nmyristoyltransferase (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 media containing NMT inhibitor or in SILAC media 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 Author Manuscript Author Manuscript

Protein acetylation describes the reversible posttranslational transfer of an acetyl group from acetyl-coenzymeA (Acetyl-CoA) to a target protein, most commonly on the side-chain amine of a lysine residue (Figure 6A). This process is catalyzed by lysine acetyltransferases (KATs) and is removed by lysine deacetylases (KDACs) (Yang et al., 2011). KDACs can be broken into two families: classic metallohydrolases that act directly to hydrolyze acetimide and the sirtuins that utilize an nitotinamide 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 one year later, Vincent Allfrey and colleagues showed that radiolabelled acetate was rapidly sequestered from media 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. Additionally, 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 had 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


Chuh et al.

Page 19

Author Manuscript

modulation of protein acetylation levels as a therapeutic strategy. For example Vorinostat, a histone deacetylase (HDAC) inhibitor, is the first FDA-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 solid phase peptide synthesis is quite straight forward. 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 (S. He et al., 2003). For example, NCL was used to generate histone H4 site specifically acetylated at K16 (ShogrenKnaak 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 6B) (Neumann et al., 2009; 2008). The M. bakeri pyrrolysyl tRNA synthetase/tRNACUA was once again used to incorporate acetylated 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 FRET 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 6C) (R. Huang et al., 2010). Specifically, Cole and co-workers 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 bromodomains, 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. Towards this goal, sodium 4-pentynoate has been developed as an Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 20


MCR lysine acetylation both in living cells and in vitro (Figure 6D). 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 6D) (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 lab introduced a chemical reporter of aspirin acetylation, AspAlk (Figure 6D) 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 (J. 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 7A) (Biggar and Li, 2015; D. Y. Lee et al., 2005). Lysine residues can be mono-, di- or tri-methylated, while arginines can monomethylated or di-methylated. Histidines have only been reported to be mono-methylated, however, this modification is uncommon and its role remains unclear (Greer and Shi, 2012). Histone methylation plays well-documented roles in transcriptional regulation, and nonhistone 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 chromatinassociating non-histone proteins (J. 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


Chuh et al.

Page 21

Author Manuscript

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 sitespecifically 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 solid phase peptide synthesis; 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 (S. 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 a series of acetylated lysine residues in histone H4 were prepared (Ruthenburg et al., 2011). Using these homogenous 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 semi-synthetic 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 7B). A pyrrolysyl-tRNA synthetase pair has been used extensively for incorporation of mono- and dimethylated lysine into recombinant proteins as it prefers methylated lysine over unmodified lysine. In one example, mono- and dimethylated 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 (D. P. 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; Y.-S. Wang et al., 2010). The Chin lab has also used a GOPAL-like strategy to reveal a specific lysine residue that can be di-methylated using reductive methylation (D. P. Nguyen et al., 2010). Unnatural amino acid mutagenesis methods are yet to be used for the direct incorporation of di- and tri-methylated lysine residues. However, a chemical method for the facile incorporation of mono-, di, and tri-methylated lysine analogs was developed (Figure 7C) (Simon et al., 2007). Briefly, cysteine resides at the desired site of modification can be selectively alkylated it to create a methyl lysine analog (MLA), Nmethylated aminoethylcysteine. Notably, these mono-, di-, and tri-methylated lysine analogs were functionally similar to their native counterparts when incorporated into recombinant proteins.


Chuh et al.

Page 22

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 phosphorylation were developed as described above. Towards this goal, an additional series of azide- and alkyne-SAM analogs of various sizes were prepared (Luo, 2012). Several of the developed SAM analogs 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 7D) (Islam et al., 2013). Although this strategy was useful for in vitro testing and the screening of cellular extract, 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 (R. Wang et al., 2013). 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 selenium-based 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

Author Manuscript

Protein adenosine diphosphate (ADP) ribosylation describes the posttranslational transfer of an ADP-ribose moiety from β-nicotinamide adenine dinucleotide (NAD+) to a variety of amino acid side chains on protein acceptors, including aspartate, glutamate, lysine, arginine and cysteine (Figure 7E) (Daniels et al., 2015; Leung, 2014). This mono-ADP-ribosylation is then often polymerized to generate a long chain of repeating units, termed poly-ADPribosylation. In humans, ADP-ribosylation is installed by a family of 17 diphtheria toxinlike ADP-ribosyltransferases (ARTDs), commonly known as poly-ADP-ribose polymerases (PARPs). 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-ADPribose 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 is 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


Chuh et al.

Page 23

Author Manuscript

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 majority of ribosylation events, in particular mono-ADP-ribosylation, is not well understood. Obstacles contributing to difficulties in studying ADP-ribosylation include 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 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.

Author Manuscript

Encoding ADP-ribosylation

Author Manuscript

As with the more extensively studied modifications above, the preparation of sitespecifically ADP-ribosylated peptides and proteins is key to investigating its biochemical effects. However, the chemically sensitive nature of the pyrophosphate bond makes peptide synthesis challenging. In order to circumvent these issues, Filippov and co-workers installed a selectively protected ribosylated asparagine or glutamine residue into peptide using solid phase peptide synthesis (van der Heden van Noort et al., 2010). An alternative approach has also been developed for the generation of ADP-ribosylation analogs (Figure 7F) (Moyle and Muir, 2010). More specifically, this method uses aminooxy-functionalized amino acids for the specific conjugation of ADP-ribose onto peptides and semi-synthetic proteins, with oxime ligation occurs between the aminooxy-functionalized amino acid of choice and the anomeric carbon of ribose at pH 4.5. Notably, this mimic of mono-ADP-ribosylated in a H2B-derived peptide was a substrate for PARP1, producing site-specific-poly-ADPribosylated peptide conjugates. Finally, the same authors used NCL to prepare ADPribosylated 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

Author Manuscript

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 analogues with cell lysate and recombinant PARP1 led to the discovery of 70 potentially new PARP1 protein substrates (Figure 7G) (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 C5’ 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 tandem mass spectrometry allowed for identification of unique targets for both enzymes, which could be further applied to all 17 Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 24

Author Manuscript

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 mono-ADP-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 sitespecific chemical reactions for the installment of modifications through either native or nonnative linkages, further expanding our toolbox. For example, in a tour-de-force of NCL mediated synthesis, the Muir lab has recently prepared histones with different combinations of ubiquitination, acetylation, and methylation marks (U. T. T. 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 mass spectrometry 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 co-workers used the Alk-16 MCR to show that changes in the palmitoylation of specific proteins controls the entry into meiosis in fission yeast, suggesting that single enzymes that install PTMs can control complex biological processes (M. M. Zhang et al., 2013).

Author Manuscript

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 labs with a 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 the 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 mass spectrometry methods, including new ionization techniques and computer programs that allow for the direct


Chuh et al.

Page 25

Author Manuscript

identification of modification sites. Chemical methods have been instrumental in the investigation of PTMs towards 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 homogenous 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.

Acknowledgments 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.

Author Manuscript

References


Abeywardana T, Lin YH, Rott R, Engelender S, Pratt MR. Site-specific differences in proteasomedependent degradation of monoubiquitinated α-synuclein. Chem Biol. 2013; 20:1207– 1213.10.1016/j.chembiol.2013.09.009 [PubMed: 24210006] Ai HW, Lee JW, Schultz PG. A method to site-specifically introduce methyllysine into proteins in E. coli. Chem Commun (Camb). 2010; 46:5506–5508.10.1039/c0cc00108b [PubMed: 20571694] Ajish Kumar KS, Haj-Yahya M, Olschewski D, Lashuel HA, Brik A. Highly efficient and chemoselective peptide ubiquitylation. Angew Chem Int Ed Engl. 2009; 48:8090–8094.10.1002/ anie.200902936 [PubMed: 19780082] Alfaro JF, Gong CX, Monroe ME, Aldrich JT, Clauss TRW, Purvine SO, Wang Z, Camp DG, Shabanowitz J, Stanley P, Hart GW, Hunt DF, Yang F, Smith RD. Tandem mass spectrometry identifies many mouse brain O-GlcNAcylated proteins including EGF domain-specific O-GlcNAc transferase targets. Proc Natl Acad Sci USA. 2012; 109:7280–7285.10.1073/pnas.1200425109 [PubMed: 22517741] Allen J, Li M, Brinkworth C, Paulson J, Wang D, Hubner A, Chou W, Davis R, Burlingame A, Messing R, Katayama C, Hedrick S, Shokat K. A semisynthetic epitope for kinase substrates. Nat Meth. 2007; 4:511–516.10.1038/nmeth1048 Allen JJ, Lazerwith SE, Shokat KM. Bio-orthogonal Affinity Purification of Direct Kinase Substrates. J Am Chem Soc. 2005; 127:5288–5289.10.1021/ja050727t [PubMed: 15826144] Allfrey VG, Faulkner R, Mirsky AE. Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc Natl Acad Sci USA. 1964; 51:786–794. [PubMed: 14172992] Alonso A, Sasin J, Bottini N, Friedberg I, Friedberg I, Osterman A, Godzik A, Hunter T, Dixon J, Mustelin T. Protein tyrosine phosphatases in the human genome. Cell. 2004; 117:699–711.10.1016/ j.cell.2004.05.018 [PubMed: 15186772] Ambler RP, Rees MW. Epsilon-N-Methyl-lysine in bacterial flagellar protein. Nature. 1959; 184:56– 57. [PubMed: 13793118] Ansaloni A, Wang ZM, Jeong JS, Ruggeri FS, Dietler G, Lashuel HA. One-Pot Semisynthesis of Exon 1 of the Huntingtin Protein: New Tools for Elucidating the Role of Posttranslational Modifications in the Pathogenesis of Huntington’s Disease. Angew Chem Int Ed. 2014; 53:1928–1933.10.1002/ anie.201307510 Bao X, Zhao Q, Yang T, Fung YME, Li XD. A Chemical Probe for Lysine Malonylation. Angew Chem Int Ed. 2013; 52:4883–4886.10.1002/anie.201300252 Bartke T, Vermeulen M, Xhemalce B, Robson SC, Mann M, Kouzarides T. Nucleosome-interacting proteins regulated by DNA and histone methylation. Cell. 2010; 143:470–484.10.1016/j.cell. 2010.10.012 [PubMed: 21029866]


Chuh et al.

Page 26


Baskin JM, Dehnert KW, Laughlin ST, Amacher SL, Bertozzi CR. Visualizing enveloping layer glycans during zebrafish early embryogenesis. Proc Natl Acad Sci USA. 2010; 107:10360– 10365.10.1073/pnas.0912081107 [PubMed: 20489181] Bateman LA, Zaro BW, Miller SM, Pratt MR. An Alkyne–Aspirin Chemical Reporter for the Detection of Aspirin-Dependent Protein Modification in Living Cells. J Am Chem Soc. 2013; 135:14568–14573.10.1021/ja408322b [PubMed: 23998633] Baumann K, Kowalczyk D, Kunz H. Total Synthesis of the Glycopeptide Recognition Domain of the P-Selectin Glycoprotein Ligand 1. Angew Chem Int Ed. 2008; 47:3445–3449.10.1002/anie. 200705762 Bello C, Farbiarz K, Möller JF, Becker CFW, Schwientek T. A quantitative and site-specific chemoenzymatic glycosylation approach for PEGylated MUC1 peptides. Chem Sci. 2014; 5:1634– 8.10.1039/c3sc52641k Berndt N, Hamilton AD, Sebti SM. Targeting protein prenylation for cancer therapy. Nat Rev Cancer. 2011; 11:775–791.10.1038/nrc3151 [PubMed: 22020205] Biggar KK, Li SSC. Non-histone protein methylation as a regulator of cellular signalling and function. Nat Rev Mol Cell Biol. 2015; 16:5–17.10.1038/nrm3915 [PubMed: 25491103] Binda O, Boyce M, Rush JS, Palaniappan KK, Bertozzi CR, Gozani O. A chemical method for labeling lysine methyltransferase substrates. ChemBioChem. 2011; 12:330–334.10.1002/cbic. 201000433 [PubMed: 21243721] Bishop AC, Shah K, Liu Y, Witucki L, Kung C, Shokat KM. Design of allele-specific inhibitors to probe protein kinase signaling. Curr Biol. 1998; 8:257–266. [PubMed: 9501066] Bishop AC, Ubersax JA, Petsch DT, Matheos DP, Gray NS, Blethrow J, Shimizu E, Tsien JZ, Schultz PG, Rose MD, Wood JL, Morgan DO, Shokat KM. A chemical switch for inhibitor-sensitive alleles of any protein kinase. Nature. 2000; 407:395–401.10.1038/35030148 [PubMed: 11014197] Blethrow JD, Glavy JS, Morgan DO, Shokat KM. Covalent capture of kinase-specific phosphopeptides reveals Cdk1-cyclin B substrates. Proc Natl Acad Sci USA. 2008; 105:1442–1447.10.1073/pnas. 0708966105 [PubMed: 18234856] Borodovsky A, Kessler B, Casagrande R, Overkleeft H, Wilkinson K, Ploegh H. A novel active sitedirected probe specific for deubiquitylating enzymes reveals proteasome association of USP14. EMBO J. 2001; 20:5187–5196. [PubMed: 11566882] Borodovsky A, Ovaa H, Kolli N, Gan-Erdene T, Wilkinson K, Ploegh H, Kessler B. Chemistry-based functional proteomics reveals novel members of the deubiquitinating enzyme family. 2002; 9:1149–1159. Bothwell IR, Islam K, Chen Y, Zheng W, Blum G, Deng H, Luo M. Se-Adenosyl- l-selenomethionine Cofactor Analogue as a Reporter of Protein Methylation. J Am Chem Soc. 2012; 134:14905– 14912.10.1021/ja304782r [PubMed: 22917021] Brunsveld L, Kuhlmann J, Waldmann H. Synthesis of palmitoylated Ras-peptides and -proteins. Methods. 2006; 40:151–165.10.1016/j.ymeth.2006.04.014 [PubMed: 17012027] Calce E, Leone M, Monfregola L, De Luca S. Lipidated peptides via post-synthetic thioalkylation promoted by molecular sieves. Amino Acids. 2014; 46:1899–1905.10.1007/s00726-014-1742-y [PubMed: 24748120] Capdeville R, Buchdunger E, Zimmermann J, Matter A. Glivec (STI571, imatinib), a rationally developed, targeted anticancer drug. Nat Rev Drug Discov. 2002; 1:493–502.10.1038/nrd839 [PubMed: 12120256] Carter-O’Connell I, Jin H, Morgan RK, David LL, Cohen MS. Engineering the substrate specificity of ADP-ribosyltransferases for identifying direct protein targets. J Am Chem Soc. 2014; 136:5201– 5204.10.1021/ja412897a [PubMed: 24641686] Cato D, Buskas T, Boons GJ. Highly Efficient Stereospecific Preparation of Tn and TF Building Blocks Using Thioglycosyl Donors and the Ph 2 SO/Tf 2 O Promotor System*. Journal of Carbohydrate Chemistry. 2005; 24:503–516.10.1081/CAR-200067091 Chalker JM, Bernardes GJL, Davis BG. A “Tag-and-Modify” Approach to Site-Selective Protein Modification. Acc Chem Res. 2011; 44:730–741.10.1021/ar200056q [PubMed: 21563755] Chamberlain LH, Shipston MJ. The physiology of protein S-acylation. Physiol Rev. 2015; 95:341– 376.10.1152/physrev.00032.2014 [PubMed: 25834228]


Chuh et al.

Page 27


Charron G, Li MMH, MacDonald MR, Hang HC. Prenylome profiling reveals S-farnesylation is crucial for membrane targeting and antiviral activity of ZAP long-isoform. Proc Natl Acad Sci USA. 2013; 110:11085–11090.10.1073/pnas.1302564110 [PubMed: 23776219] Charron G, Tsou LK, Maguire W, Yount JS, Hang HC. Alkynyl-farnesol reporters for detection of protein S-prenylation in cells. Mol Biosyst. 2011; 7:67–73.10.1039/c0mb00183j [PubMed: 21107478] Chatterjee C, McGinty RK, Fierz B, Muir TW. Disulfide-directed histone ubiquitylation reveals plasticity in hDot1L activation. Nat Chem Biol. 2010; 6:267–269.10.1038/nchembio.315 [PubMed: 20208522] Chen J, Ai Y, Wang J, Haracska L, Zhuang Z. Chemically ubiquitylated PCNA as a probe for eukaryotic translesion DNA synthesis. Nat Chem Biol. 2010; 6:270–272.10.1038/nchembio.316 [PubMed: 20208521] Chen ZJ, Sun LJ. Nonproteolytic functions of ubiquitin in cell signaling. Mol Cell. 2009; 33:275– 286.10.1016/j.molcel.2009.01.014 [PubMed: 19217402] Chi P, Allis CD, Wang GG. Covalent histone modifications–miswritten, misinterpreted and mis-erased in human cancers. Nat Rev Cancer. 2010; 10:457–469.10.1038/nrc2876 [PubMed: 20574448] Chin JW. Expanding and Reprogramming the Genetic Code of Cells and Animals. Annu Rev Biochem. 2014; 83:379–408.10.1146/annurev-biochem-060713-035737 [PubMed: 24555827] Chuh KN, Pratt MR. Chemical methods for the proteome-wide identification of posttranslationally modified proteins. Curr Opin Chem Biol. 2015a; 24:27–37.10.1016/j.cbpa.2014.10.020 [PubMed: 25461721] Chuh KN, Pratt MR. Chemistry-enabled methods for the visualization of cell-surface glycoproteins in Metazoans. Glycoconj J. 2015b; 32:443–454.10.1007/s10719-015-9589-3 [PubMed: 25913724] Chuh KN, Zaro BW, Piller F, Piller V, Pratt MR. Changes in metabolic chemical reporter structure yield a selective probe of O-GlcNAc modification. J Am Chem Soc. 2014; 136:12283– 12295.10.1021/ja504063c [PubMed: 25153642] Clark PM, Dweck JF, Mason DE, Hart CR, Buck SB, Peters EC, Agnew BJ, Hsieh-Wilson LC. Direct In-Gel Fluorescence Detection and Cellular Imaging of O-GlcNAc-Modified Proteins. J Am Chem Soc. 2008; 130:11576–11577.10.1021/ja8030467 [PubMed: 18683930] Cole CM, Yang J, Šečkutė J, Devaraj NK. Fluorescent Live-Cell Imaging of Metabolically Incorporated Unnatural Cyclopropene-Mannosamine Derivatives. ChemBioChem. 2013; 14:205– 208.10.1002/cbic.201200719 [PubMed: 23292753] Curtin NJ, Szabo C. Therapeutic applications of PARP inhibitors: Anticancer therapy and beyond. Molecular Aspects of Medicine. 2013; 34:1217–1256.10.1016/j.mam.2013.01.006 [PubMed: 23370117] Dancy BCR, Ming SA, Papazyan R, Jelinek CA, Majumdar A, Sun Y, Dancy BM, Drury WJ III, Cotter RJ, Taverna SD, Cole PA. Azalysine Analogues as Probes for Protein Lysine Deacetylation and Demethylation. J Am Chem Soc. 2012; 12031215234000110.1021/ja209574z Daniels CM, Ong SE, Leung AKL. The Promise of Proteomics for the Study of ADP-Ribosylation. Mol Cell. 2015; 58:911–924.10.1016/j.molcel.2015.06.012 [PubMed: 26091340] Davis B. Synthesis of glycoproteins. Chem Rev. 2002; 102:579–601. [PubMed: 11841255] Davis L, Chin JW. Designer proteins: applications of genetic code expansion in cell biology. Nat Rev Mol Cell Biol. 2012; 13:168–182.10.1038/nrm3286 [PubMed: 22334143] Dawson P, Muir T, Clark-Lewis I, Kent S. Synthesis of proteins by native chemical ligation. Science. 1994; 266:776–779. [PubMed: 7973629] DeGraw AJ, Palsuledesai C, Ochocki JD, Dozier JK, Lenevich S, Rashidian M, Distefano MD. Evaluation of alkyne-modified isoprenoids as chemical reporters of protein prenylation. Chem Biol Drug Des. 2010; 76:460–471.10.1111/j.1747-0285.2010.01037.x [PubMed: 21040496] Dias W, Hart G. O-GlcNAc modification in diabetes and Alzheimer’s disease. Mol BioSys. 2007; 3:766–772. Dondoni A, Massi A, Nanni P, Roda A. A new ligation strategy for peptide and protein glycosylation: photoinduced thiol-ene coupling. Chemistry-A European Journal. 2009; 15:11444–11449.10.1002/ chem.200901746


Chuh et al.

Page 28


Doores KJ, Mimura Y, Dwek RA, Rudd PM, Elliott T, Davis BG. Direct deprotected glycosylasparagine ligation. Chem Commun. 2006:1401–1403.10.1039/b515472c Drisdel RC, Green WN. Labeling and quantifying sites of protein palmitoylation. BioTechniques. 2004; 36:276–285. [PubMed: 14989092] Du J, Jiang H, Lin H. Investigating the ADP-ribosyltransferase activity of sirtuins with NAD analogues and 32P-NAD. Biochemistry. 2009; 48:2878–2890.10.1021/bi802093g [PubMed: 19220062] Eger S, Castrec B, Hübscher U, Scheffner M, Rubini M, Marx A. Generation of a Monoubiquitinated PCNA Mimic by Click Chemistry. ChemBioChem. 2011; 12:2807–2812.10.1002/cbic.201100444 [PubMed: 22052741] Ekkebus R, van Kasteren SI, Kulathu Y, Scholten A, Berlin I, Geurink PP, De Jong A, Goerdayal S, Neefjes J, Heck AJR, Komander D, Ovaa H. On Terminal Alkynes That Can React with ActiveSite Cysteine Nucleophiles in Proteases. J Am Chem Soc. 2013; 135:2867–2870.10.1021/ ja309802n [PubMed: 23387960] Fernández-González M, Boutureira O, Bernardes GJL, Chalker JM, Young MA, Errey JC, Davis BG. Site-selective chemoenzymatic construction of synthetic glycoproteins using endoglycosidases. Chem Sci. 2010; 1:709–7.10.1039/c0sc00265h Floyd N, Vijayakrishnan B, Koeppe JR, Davis BG. Thiyl glycosylation of olefinic proteins: S-linked glycoconjugate synthesis. Angew Chem Int Ed Engl. 2009; 48:7798–7802.10.1002/anie. 200903135 [PubMed: 19739166] Fröhlich M, Dejanovic B, Kashkar H, Schwarz G, Nussberger S. S-palmitoylation represents a novel mechanism regulating the mitochondrial targeting of BAX and initiation of apoptosis. Cell Death Dis. 2014; 5:e1057.10.1038/cddis.2014.17 [PubMed: 24525733] Gamblin D, Scanlan E, Davis B. Glycoprotein synthesis: an update. Chem Rev. 2009; 109:131–163. doi:10.1021/cr078291i [pii]. [PubMed: 19093879] Gamblin DP, van Kasteren S, Bernardes GJL, Chalker JM, Oldham NJ, Fairbanks AJ, Davis BG. Chemical site-selective prenylation of proteins. Mol Biosyst. 2008; 4:558–561.10.1039/b802199f [PubMed: 18493653] Garber KCA, Carlson EE. Thiol-ene enabled detection of thiophosphorylated kinase substrates. ACS Chem Biol. 2013; 8:1671–1676.10.1021/cb400184v [PubMed: 23668631] Goodfellow JJ, Baruah K, Yamamoto K, Bonomelli C, Krishna B, Harvey DJ, Crispin M, Scanlan CN, Davis BG. An Endoglycosidase with Alternative Glycan Specificity Allows Broadened Glycoprotein Remodelling. J Am Chem Soc. 2012; 134:8030–8033.10.1021/ja301334b [PubMed: 22551167] Grammel M, Hang HC. chemical reporters for biological discovery. Nat Chem Biol. 2013; 9:475– 484.10.1038/nchembio.1296 [PubMed: 23868317] Greer EL, Shi Y. Histone methylation: a dynamic mark in health, disease and inheritance. Nat Rev Genet. 2012; 13:343–357.10.1038/nrg3173 [PubMed: 22473383] Groff D, Chen PR, Peters FB, Schultz PG. A Genetically Encoded ε-N-Methyl Lysine in Mammalian Cells. ChemBioChem. 2010; 11:1066–1068.10.1002/cbic.200900690 [PubMed: 20422671] Grogan M, Pratt M, Marcaurelle L, Bertozzi C. Homogeneous Glycopeptides and Glycoproteins for Biological Investigation. Ann Rev Biochem. 2002; 71:593–634. [PubMed: 12045107] Gross S, Rahal R, Stransky N, Lengauer C, Hoeflich KP. Targeting cancer with kinase inhibitors. J Clin Invest. 2015; 125:1780–1789.10.1172/JCI76094 [PubMed: 25932675] Hahn ME, Muir TW. Photocontrol of Smad2, a Multiphosphorylated Cell-Signaling Protein, through Caging of Activating Phosphoserines. Angew Chem Int Ed Engl. 2004; 43:5800–5803.10.1002/ anie.200461141 [PubMed: 15523718] Haj-Yahya M, Fauvet B, Herman-Bachinsky Y, Hejjaoui M, Bavikar SN, Karthikeyan SV, Ciechanover A, Lashuel HA, Brik A. Synthetic polyubiquitinated α-Synuclein reveals important insights into the roles of the ubiquitin chain in regulating its pathophysiology. Proc Natl Acad Sci USA. 2013; 110:17726–17731.10.1073/pnas.1315654110 [PubMed: 24043770] Haneda K, Inazu T, Mizuno M, Iguchi R, Yamamoto K, Kumagai H, Aimoto S, Suzuki H, Noda T. Chemo-enzymatic synthesis of calcitonin derivatives containing N-linked oligosaccharides. Bioorg Med Chem Lett. 1998; 8:1303–1306. [PubMed: 9871755]


Chuh et al.

Page 29


Hang HC, Bertozzi CR. The chemistry and biology of mucin-type O-linked glycosylation. Bioorg Med Chem. 2005; 13:5021–5034.10.1016/j.bmc.2005.04.085 [PubMed: 16005634] Hang HC, Linder ME. Exploring Protein Lipidation with Chemical Biology. Chem Rev. 2011; 111:6341–6358.10.1021/cr2001977 [PubMed: 21919527] Hang HC, Wilson JP, Charron G. Bioorthogonal Chemical Reporters for Analyzing Protein Lipidation and Lipid Trafficking. Acc Chem Res. 2011; 44:699–708.10.1021/ar200063v [PubMed: 21675729] Hang HC, Yu C, Kato DL, Bertozzi CR. A metabolic labeling approach toward proteomic analysis of mucin-type O-linked glycosylation. Proc Natl Acad Sci USA. 2003; 100:14846–14851.10.1073/ pnas.2335201100 [PubMed: 14657396] Hannoush RN. Synthetic protein lipidation. Curr Opin Chem Biol. 2015; 28:39–46.10.1016/j.cbpa. 2015.05.025 [PubMed: 26080277] Hayes JS, Mayer SE. Regulation of guinea pig heart phosphorylase kinase by cAMP, protein kinase, and calcium. Am J Physiol. 1981; 240:E340–9. [PubMed: 6259950] He S, Bauman D, Davis JS, Loyola A, Nishioka K, Gronlund JL, Reinberg D, Meng F, Kelleher N, McCafferty DG. Facile synthesis of site-specifically acetylated and methylated histone proteins: reagents for evaluation of the histone code hypothesis. Proc Natl Acad Sci USA. 2003; 100:12033–12038.10.1073/pnas.2035256100 [PubMed: 14530408] He Y, Hinklin R, Chang J, Kiessling L. Stereoselective N-glycosylation by Staudinger ligation. Org Lett. 2004; 6:4479–4482. [PubMed: 15548055] Heal WP, Wickramasinghe SR, Leatherbarrow RJ, Tate EW. N-Myristoyl transferase-mediated protein labelling in vivo. Org Biomol Chem. 2008; 6:2308–2315.10.1039/b803258k [PubMed: 18563263] Helenius A, Aebi M. Intracellular Functions of N-linked Glycans. Science. 2001; 291:2364–2369. [PubMed: 11269317] Hemantha HP, Bavikar SN, Herman-Bachinsky Y, Haj-Yahya N, Bondalapati S, Ciechanover A, Brik A. Nonenzymatic Polyubiquitination of Expressed Proteins. J Am Chem Soc. 2014; 136:2665– 2673.10.1021/ja412594d [PubMed: 24437386] Hershko A, Ciechanover A. The ubiquitin system. Annu Rev Biochem. 1998; 67:425–479. [PubMed: 9759494] Hsu TL, Hanson SR, Kishikawa K, Wang SK, Sawa M, Wong CH. Alkynyl sugar analogs for the labeling and visualization of glycoconjugates in cells. Proc Natl Acad Sci USA. 2007; 104:2614– 2619.10.1073/pnas.0611307104 [PubMed: 17296930] Huang J, Berger SL. The emerging field of dynamic lysine methylation of non-histone proteins. Curr Opin Genet Dev. 2008; 18:152–158.10.1016/j.gde.2008.01.012 [PubMed: 18339539] Huang R, Holbert MA, Tarrant MK, Curtet S, Colquhoun DR, Dancy BM, Dancy BC, Hwang Y, Tang Y, Meeth K, Marmorstein R, Cole RN, Khochbin S, Cole PA. Site-Specific Introduction of an Acetyl-Lysine Mimic into Peptides and Proteins by Cysteine Alkylation. J Am Chem Soc. 2010; 132:9986–9987.10.1021/ja103954u [PubMed: 20608637] Huang W, Li J, Wang LX. Unusual transglycosylation activity of Flavobacterium meningosepticum endoglycosidases enables convergent chemoenzymatic synthesis of core fucosylated complex Nglycopeptides. ChemBioChem. 2011; 12:932–941.10.1002/cbic.201000763 [PubMed: 21374780] Huse M, Muir T, Xu L, Chen Y, Kuriyan J, Massague J. The TGF beta receptor activation process: an inhibitor- to substrate-binding switch. Mol Cell. 2001; 8:671–682. [PubMed: 11583628] Islam K, Chen Y, Wu H, Bothwell IR, Blum GJ, Zeng H, Dong A, Zheng W, Min J, Deng H, Luo M. Defining efficient enzyme-cofactor pairs for bioorthogonal profiling of protein methylation. Proc Natl Acad Sci USA. 2013; 110:16778–16783.10.1073/pnas.1216365110 [PubMed: 24082136] Ivaldi C, Martin BR, Kieffer-Jaquinod S, Chapel A, Levade T, Garin J, Journet A. Proteomic analysis of S-acylated proteins in human B cells reveals palmitoylation of the immune regulators CD20 and CD23. PLoS ONE. 2012; 7:e37187.10.1371/journal.pone.0037187 [PubMed: 22615937] Iwase, S.; Shi, Y. Histone and DNA Modifications in Mental Retardation. Springer Basel; Basel: 2010. Histone and DNA Modifications in Mental Retardation; p. 147-173. Izumi M, Makimura Y, Dedola S, Seko A, Kanamori A, Sakono M, Ito Y, Kajihara Y. Chemical Synthesis of Intentionally Misfolded Homogeneous Glycoprotein: A Unique Approach for the


Chuh et al.

Page 30


Study of Glycoprotein Quality Control. J Am Chem Soc. 2012; 134:7238–7241.10.1021/ ja3013177 [PubMed: 22497239] Jiang H, Kim JH, Frizzell KM, Kraus WL, Lin H. Clickable NAD analogues for labeling substrate proteins of poly(ADP-ribose) polymerases. J Am Chem Soc. 2010; 132:9363–9372.10.1021/ ja101588r [PubMed: 20560583] Jones ML, Collins MO, Goulding D, Choudhary JS, Rayner JC. Analysis of protein palmitoylation reveals a pervasive role in Plasmodium development and pathogenesis. Cell Host Microbe. 2012; 12:246–258.10.1016/j.chom.2012.06.005 [PubMed: 22901544] Khidekel N, Ficarro S, Clark P, Bryan M, Swaney D, Rexach J, Sun Y, Coon J, Peters E, HsiehWilson L. Probing the dynamics of O-GlcNAc glycosylation in the brain using quantitative proteomics. Nat Chem Biol. 2007; 3:339–348. [PubMed: 17496889] Kim S, Paik WK. Studies on the Origin of e-N-Methyl-L-lysine in Protein*. J Biol Chem. 1965; 240:4629–4634. [PubMed: 5846984] Komander D, Clague MJ, Urbé S. Breaking the chains: structure and function of the deubiquitinases. Nat Rev Mol Cell Biol. 2009; 10:550–563.10.1038/nrm2731 [PubMed: 19626045] Komander D, Rape M. The Ubiquitin Code. Annu Rev Biochem. 2012; 81:203–229.10.1146/annurevbiochem-060310-170328 [PubMed: 22524316] Kouzarides T. Chromatin modifications and their function. Cell. 2007; 128:693–705.10.1016/j.cell. 2007.02.005 [PubMed: 17320507] Kragol G, Lumbierres M, Palomo JM, Waldmann H. Solid-Phase Synthesis of Lipidated Peptides. Angew Chem Int Ed. 2004; 43:5839–5842.10.1002/anie.200461150 Kulathu Y, Komander D. Atypical ubiquitylation — the unexplored world of polyubiquitin beyond Lys48 and Lys63 linkages. Nat Rev Mol Cell Biol. 2012; 13:508–523.10.1038/nrm3394 [PubMed: 22820888] Kumar KSA, Spasser L, Erlich LA, Bavikar SN, Brik A. Total chemical synthesis of diubiquitin chains. Angew Chem Int Ed Engl. 2010; 49:9126–9131.10.1002/anie.201003763 [PubMed: 20815002] Lang K, Chin JW. Cellular incorporation of unnatural amino acids and bioorthogonal labeling of proteins. Chem Rev. 2014; 114:4764–4806.10.1021/cr400355w [PubMed: 24655057] Laughlin S, Baskin J, Amacher S, Bertozzi C. In vivo imaging of membrane-associated glycans in developing zebrafish. Science. 2008; 320:664–667. doi:320/5876/664 [pii] 10.1126/science. 1155106. [PubMed: 18451302] Laughlin ST, Bertozzi CR. In Vivo Imaging of Caenorhabditis elegans Glycans. ACS Chem Biol. 2009; 4:1068–1072.10.1021/cb900254y [PubMed: 19954190] Lee DY, Teyssier C, Strahl BD, Stallcup MR. Role of Protein Methylation in Regulation of Transcription. Endocrine Reviews. 2005; 26:147–170.10.1210/er.2004-0008 [PubMed: 15479858] Lee S, Oh S, Yang A, Kim J, Söll D, Lee D, Park HS. A facile strategy for selective incorporation of phosphoserine into histones. Angew Chem Int Ed Engl. 2013; 52:5771–5775.10.1002/anie. 201300531 [PubMed: 23533151] Leung AKL. Poly(ADP-ribose): an organizer of cellular architecture. J Cell Biol. 2014; 205:613– 619.10.1083/jcb.201402114 [PubMed: 24914234] Liu CC, Schultz PG. Adding new chemistries to the genetic code. Annu Rev Biochem. 2010; 79:413– 444.10.1146/annurev.biochem.052308.105824 [PubMed: 20307192] Liu Y, Shah K, Yang F, Witucki L, Shokat KM. Engineering Src family protein kinases with unnatural nucleotide specificity. Chem Biol. 1998; 5:91–101. [PubMed: 9495830] Love KR, Pandya RK, Spooner E, Ploegh HL. Ubiquitin C-terminal electrophiles are activity-based probes for identification and mechanistic study of ubiquitin conjugating machinery. ACS Chem Biol. 2009; 4:275–287.10.1021/cb9000348 [PubMed: 19256548] Lu W, Gong D, Bar-Sagi D, Cole PA. Site-specific incorporation of a phosphotyrosine mimetic reveals a role for tyrosine phosphorylation of SHP-2 in cell signaling. Mol Cell. 2001; 8:759–769. [PubMed: 11684012] Lu W, Shen K, Cole PA. Chemical dissection of the effects of tyrosine phosphorylation of SHP-2. Biochemistry. 2003; 42:5461–5468.10.1021/bi0340144 [PubMed: 12731888] Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 31


Ludolph B, Eisele F, Waldmann H. Solid-Phase Synthesis of Lipidated Peptides. J Am Chem Soc. 2002; 124:5954–5955.10.1021/ja025768t [PubMed: 12022824] Lumbierres M, Palomo JM, Kragol G, Roehrs S, Müller O, Waldmann H. Solid-Phase Synthesis of Lipidated Peptides. Chem Eur J. 2005; 11:7405–7415.10.1002/chem.200500476 [PubMed: 16193522] Luo M. Current chemical biology approaches to interrogate protein methyltransferases. ACS Chem Biol. 2012; 7:443–463.10.1021/cb200519y [PubMed: 22220966] Ma Z, Vosseller K. O-GlcNAc in cancer biology. Amino Acids. 2013; 45:719–733.10.1007/ s00726-013-1543-8 [PubMed: 23836420] Maly D, Allen J, Shokat K. A mechanism-based cross-linker for the identification of kinase-substrate pairs. J Am Chem Soc. 2004; 126:9160–9161. [PubMed: 15281787] Manning G, Whyte DB, Martinez R, Hunter T, Sudarsanam S. The protein kinase complement of the human genome. Science. 2002; 298:1912–1934.10.1126/science.1075762 [PubMed: 12471243] Marcaurelle L, Bertozzi C. Chemoselective Elaboration of O-Linked Glycopeptide Mimetics by Alkylation of 3-ThioGalNAc. J Am Chem Soc. 2001; 123:1587–1595. [PubMed: 11456757] Marotta NP, Lin YH, Lewis YE, Ambroso MR, Zaro BW, Roth MT, Arnold DB, Langen R, Pratt MR. O-GlcNAc modification blocks the aggregation and toxicity of the protein α-synuclein associated with Parkinson’s disease. Nature Chemistry. 2015:1–8.10.1038/nchem.2361 Martin BR, Wang C, Adibekian A, Tully SE, Cravatt BF. Global profiling of dynamic protein palmitoylation. Nat Meth. 2012; 9:84–89.10.1038/nmeth.1769 Martin DDO, Beauchamp E, Berthiaume LG. Post-translational myristoylation: Fat matters in cellular life and death. Biochimie. 2011; 93:18–31.10.1016/j.biochi.2010.10.018 [PubMed: 21056615] Massagué J. TGFβ signalling in context. Nat Rev Mol Cell Biol. 2012; 13:616–630.10.1038/nrm3434 [PubMed: 22992590] McGinty RK, Kim J, Chatterjee C, Roeder RG, Muir TW. Chemically ubiquitylated histone H2B stimulates hDot1L-mediated intranucleosomal methylation. Nature. 2008; 453:812–816.10.1038/ nature06906 [PubMed: 18449190] Meier F, Abeywardana T, Dhall A, Marotta NP, Varkey J, Langen R, Chatterjee C, Pratt MR. Semisynthetic, Site-Specific Ubiquitin Modification of α-Synuclein Reveals Differential Effects on Aggregation. J Am Chem Soc. 2012; 134:5468–5471.10.1021/ja300094r [PubMed: 22404520] Morgan RK, Cohen MS. A Clickable Aminooxy Probe for Monitoring Cellular ADP-Ribosylation. ACS Chem Biol. 2015; 10:1778–1784.10.1021/acschembio.5b00213 [PubMed: 25978521] Moyle PM, Muir TW. Method for the synthesis of mono-ADP-ribose conjugated peptides. J Am Chem Soc. 2010; 132:15878–15880.10.1021/ja1064312 [PubMed: 20968292] Muir TW. Semisynthesis of proteins by expressed protein ligation. Annu Rev Biochem. 2003; 72:249– 289.10.1146/annurev.biochem.72.121801.161900 [PubMed: 12626339] Muir TW, Sondhi D, Cole PA. Expressed protein ligation: a general method for protein engineering. Proc Natl Acad Sci USA. 1998; 95:6705–6710. [PubMed: 9618476] Murray K. The occurrence of iε-N-methyl lysine in histones. Biochemistry. 1964; 3:10–15.10.1021/ bi00889a003 [PubMed: 14114491] Nägele E, Schelhaas M, Kuder N, Waldmann H. Chemoenzymatic synthesis of N-Ras lipopeptides. J Am Chem Soc. 1998; 120:6889–6902.10.1021/ja9805627 Neumann H, Hancock SM, Buning R, Routh A, Chapman L, Somers J, Owen-Hughes T, van Noort J, Rhodes D, Chin JW. A Method for Genetically Installing Site-Specific Acetylation in Recombinant Histones Defines the Effects of H3 K56 Acetylation. Mol Cell. 2009; 36:153– 163.10.1016/j.molcel.2009.07.027 [PubMed: 19818718] Neumann H, Peak-Chew SY, Chin JW. Genetically encoding N(epsilon)-acetyllysine in recombinant proteins. Nat Chem Biol. 2008; 4:232–234.10.1038/nchembio.73 [PubMed: 18278036] Nguyen DP, Garcia Alai MM, Kapadnis PB, Neumann H, Chin JW. Genetically encoding N(epsilon)methyl-L-lysine in recombinant histones. J Am Chem Soc. 2009; 131:14194–14195.10.1021/ ja906603s [PubMed: 19772323]


Chuh et al.

Page 32


Nguyen DP, Garcia Alai MM, Virdee S, Chin JW. Genetically directing ε-N, N-dimethyl-L-lysine in recombinant histones. Chem Biol. 2010; 17:1072–1076.10.1016/j.chembiol.2010.07.013 [PubMed: 21035729] Nguyen UTT, Bittova L, Müller MM, Fierz B, David Y, Houck-Loomis B, Feng V, Dann GP, Muir TW. Accelerated chromatin biochemistry using DNA-barcoded nucleosome libraries. Nat Meth. 2014; 11:834–840.10.1038/nmeth.3022 Niederwieser A, Späte AK, Nguyen LD, Jüngst C, Reutter W, Wittmann V. Two-Color Glycan Labeling of Live Cells by a Combination of Diels-Alder and Click Chemistry. Angew Chem Int Ed. 2013; 52:4265–4268.10.1002/anie.201208991 Nomura DK, Dix MM, Cravatt BF. Activity-based protein profiling for biochemical pathway discovery in cancer. Nat Rev Cancer. 2010; 10:630–638.10.1038/nrc2901 [PubMed: 20703252] Ortiz-Meoz RF, Merbl Y, Kirschner MW, Walker S. Microarray discovery of new OGT substrates: the medulloblastoma oncogene OTX2 is O-GlcNAcylated. J Am Chem Soc. 2014; 136:4845– 4848.10.1021/ja500451w [PubMed: 24580054] Ottesen JJ, Huse M, Sekedat MD, Muir TW. Semisynthesis of phosphovariants of Smad2 reveals a substrate preference of the activated T beta RI kinase. Biochemistry. 2004; 43:5698– 5706.10.1021/bi0498407 [PubMed: 15134444] El Oualid F, Merkx R, Ekkebus R, Hameed DS, Smit JJ, De Jong A, Hilkmann H, Sixma TK, Ovaa H. Chemical Synthesis of Ubiquitin, Ubiquitin-Based Probes, and Diubiquitin. Angew Chem Int Ed. 2010; 49:10149–10153.10.1002/anie.201005995 Pachamuthu K, Zhu X, Schmidt RR. Reversed approach to S-farnesylation and S-palmitoylation: application to an efficient synthesis of the C-terminus of lipidated human N-ras hexapeptide. The Journal of Organic Chemistry. 2005; 70:3720–3723.10.1021/jo0482357 [PubMed: 15845014] Pan Z, Scheerens H, Li SJ, Schultz BE, Sprengeler PA, Burrill LC, Mendonca RV, Sweeney MD, Scott KCK, Grothaus PG, Jeffery DA, Spoerke JM, Honigberg LA, Young PR, Dalrymple SA, Palmer JT. Discovery of selective irreversible inhibitors for Bruton’s tyrosine kinase. ChemMedChem. 2007; 2:58–61.10.1002/cmdc.200600221 [PubMed: 17154430] Parang K, Kohn JA, Saldanha SA, Cole PA. Development of photo-crosslinking reagents for protein kinase-substrate interactions. FEBS Lett. 2002; 520:156–160. [PubMed: 12044889] Park HS, Hohn MJ, Umehara T, Guo LT, Osborne EM, Benner J, Noren CJ, Rinehart J, Soll D. Expanding the Genetic Code of Escherichia coli with Phosphoserine. Science. 2011; 333:1151– 1154.10.1126/science.1207203 [PubMed: 21868676] Patterson DM, Jones KA, Prescher JA. Improved cyclopropene reporters for probing protein glycosylation. Mol Biosyst. 2014; 10:1693.10.1039/c4mb00092g [PubMed: 24623192] Patterson DM, Nazarova LA, Xie B, Kamber DN, Prescher JA. Functionalized Cyclopropenes As Bioorthogonal Chemical Reporters. J Am Chem Soc. 2012; 134:18638–18643.10.1021/ ja3060436 [PubMed: 23072583] Pellois JP, Hahn ME, Muir TW. Simultaneous Triggering of Protein Activity and Fluorescence. J Am Chem Soc. 2004; 126:7170–7171.10.1021/ja0499142 [PubMed: 15186142] Peng T, Hang HC. Bifunctional fatty acid chemical reporter for analyzing S-palmitoylated membrane protein-protein interactions in mammalian cells. J Am Chem Soc. 2015; 137:556–559.10.1021/ ja502109n [PubMed: 25575299] Peters W, Willnow S, Duisken M, Kleine H, Macherey T, Duncan KE, Litchfield DW, Lüscher B, Weinhold E. Enzymatic Site-Specific Functionalization of Protein Methyltransferase Substrates with Alkynes for Click Labeling. Angew Chem Int Ed Engl. 201010.1002/anie.201001240 Phillips DM. The presence of acetyl groups of histones. Biochem J. 1963; 87:258–263. [PubMed: 13943142] Prada PO, Saad MJ. Tyrosine kinase inhibitors as novel drugs for the treatment of diabetes. Expert Opin Investig Drugs. 2013; 22:751–763.10.1517/13543784.2013.802768 Pratt MR, Bertozzi CR. Chemoselective ligation applied to the synthesis of a biantennary N-linked glycoform of CD52. J Am Chem Soc. 2003; 125:6149–6159.10.1021/ja029346v [PubMed: 12785846] Resh MD. Trafficking and signaling by fatty-acylated and prenylated proteins. Nat Chem Biol. 2006; 2:584–590.10.1038/nchembio834 [PubMed: 17051234]


Chuh et al.

Page 33


Rexach JE, Rogers CJ, Yu SH, Tao J, Sun YE, Hsieh-Wilson LC. Quantification of O-glycosylation stoichiometry and dynamics using resolvable mass tags. Nat Chem Biol. 2010; 6:645– 651.10.1038/nchembio.412 [PubMed: 20657584] Riel-Mehan, MM.; Shokat, KM. A Crosslinker Based on a Tethered Electrophile for Mapping KinaseSubstrate Networks. 2014. Rocks O, Gerauer M, Vartak N, Koch S, Huang ZP, Pechlivanis M, Kuhlmann J, Brunsveld L, Chandra A, Ellinger B, Waldmann H, Bastiaens PIH. The palmitoylation machinery is a spatially organizing system for peripheral membrane proteins. Cell. 2010; 141:458–471.10.1016/j.cell. 2010.04.007 [PubMed: 20416930] Rodriguez E, Winans K, King D, Bertozzi C. A strategy for the chemoselective synthesis of O-linked glycopeptides with native sugar-peptide linkages. J Am Chem Soc. 1997; 119:9905–9906. Rogerson DT, Sachdeva A, Wang K, Haq T, Kazlauskaite A, Hancock SM, Huguenin-Dezot N, Muqit MMK, Fry AM, Bayliss R, Chin JW. Efficient genetic encoding of phosphoserine and its nonhydrolyzable analog. Nat Chem Biol. 2015; 11:496–503.10.1038/nchembio.1823 [PubMed: 26030730] Rossin A, Durivault J, Chakhtoura-Feghali T, Lounnas N, Gagnoux-Palacios L, Hueber AO. Fas palmitoylation by the palmitoyl acyltransferase DHHC7 regulates Fas stability. Cell Death and Differentiation. 2015; 22:643–653.10.1038/cdd.2014.153 [PubMed: 25301068] Roth AF, Wan J, Bailey AO, Sun B, Kuchar JA, Green WN, Phinney BS, Yates JR, Davis NG. Global analysis of protein palmitoylation in yeast. Cell. 2006; 125:1003–1013.10.1016/j.cell. 2006.03.042 [PubMed: 16751107] Rösner D, Schneider T, Schneider D, Scheffner M, Marx A. Click chemistry for targeted protein ubiquitylation and ubiquitin chain formation. Nat Protoc. 2015; 10:1594–1611.10.1038/nprot. 2015.106 [PubMed: 26401915] Ruthenburg AJ, Li H, Milne TA, Dewell S, McGinty RK, Yuen M, Ueberheide B, Dou Y, Muir TW, Patel DJ, Allis CD. Recognition of a mononucleosomal histone modification pattern by BPTF via multivalent interactions. Cell. 2011; 145:692–706.10.1016/j.cell.2011.03.053 [PubMed: 21596426] Saxon E, Bertozzi C. Cell surface engineering by a modified Staudinger reaction. Science. 2000; 287:2007–2010. [PubMed: 10720325] Scaffidi P, Misteli T. Lamin A-dependent nuclear defects in human aging. Science. 2006; 312:1059– 1063.10.1126/science.1127168 [PubMed: 16645051] Schmittberger T, Waldmann H. Synthesis of the palmitoylated and prenylated C-terminal lipopeptides of the human R- and N-Ras proteins. Bioorg Med Chem. 1999; 7:749–762. [PubMed: 10400328] Schreiber V, Dantzer F, Ame JC, de Murcia G. Poly(ADP-ribose): novel functions for an old molecule. Nature Publishing Group. 2006; 7:517–528.10.1038/nrm1963 Schwarzer D, Zhang Z, Zheng W, Cole PA. Negative regulation of a protein tyrosine phosphatase by tyrosine phosphorylation. J Am Chem Soc. 2006; 128:4192–4193.10.1021/ja0585174 [PubMed: 16568970] Shah K, Liu Y, Deirmengian C, Shokat KM. Engineering unnatural nucleotide specificity for Rous sarcoma virus tyrosine kinase to uniquely label its direct substrates. Proc Natl Acad Sci USA. 1997; 94:3565–3570. [PubMed: 9108016] Shah NH, Dann GP, Vila-Perelló M, Liu Z, Muir TW. Ultrafast Protein Splicing is Common among Cyanobacterial Split Inteins: Implications for Protein Engineering. J Am Chem Soc. 2012; 134:11338–11341.10.1021/ja303226x [PubMed: 22734434] Shah NH, Muir TW. Inteins: nature’s gift to protein chemists. Chem Sci. 2013; 5:446.10.1039/ c3sc52951g [PubMed: 24634716] Shahbazian MD, Grunstein M. Functions of site-specific histone acetylation and deacetylation. Annu Rev Biochem. 2007; 76:75–100.10.1146/annurev.biochem.76.052705.162114 [PubMed: 17362198] Shanmugham A, Fish A, Luna-Vargas MPA, Faesen AC, El Oualid F, Sixma TK, Ovaa H. Nonhydrolyzable ubiquitin-isopeptide isosteres as deubiquitinating enzyme probes. J Am Chem Soc. 2010; 132:8834–8835.10.1021/ja101803s [PubMed: 20540574]


Chuh et al.

Page 34


Shirakawa K, Chavez L, Hakre S, Calvanese V, Verdin E. Reactivation of latent HIV by histone deacetylase inhibitors. Trends Microbiol. 2013; 21:277–285.10.1016/j.tim.2013.02.005 [PubMed: 23517573] Shogren-Knaak M, Ishii H, Sun JM, Pazin MJ, Davie JR, Peterson CL. Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science. 2006; 311:844–847.10.1126/ science.1124000 [PubMed: 16469925] Simon MD, Chu F, Racki LR, de la Cruz CC, Burlingame AL, Panning B, Narlikar GJ, Shokat KM. The site-specific installation of methyl-lysine analogs into recombinant histones. Cell. 2007; 128:1003–1012.10.1016/j.cell.2006.12.041 [PubMed: 17350582] Slade PG, Hajivandi M, Bartel CM, Gorfien SF. Identifying the CHO secretome using mucin-type Olinked glycosylation and click-chemistry. J Proteome Res. 2012; 11:6175–6186.10.1021/ pr300810f [PubMed: 23140450] Sommer S, Weikart ND, Brockmeyer A, Janning P, Mootz HD. Expanded Click Conjugation of Recombinant Proteins with Ubiquitin-Like Modifiers Reveals Altered Substrate Preference of SUMO2-Modified Ubc9. Angew Chem Int Ed. 2011; 50:9888–9892.10.1002/anie.201102531 Späte AK, Bußkamp H, Niederwieser A, Schart VF, Marx A, Wittmann V. Rapid Labeling of Metabolically Engineered Cell-Surface Glycoconjugates with a Carbamate-Linked Cyclopropene Reporter. Bioconjugate Chem. 2014; 25:147–154.10.1021/bc4004487 Statsuk AV, Maly DJ, Seeliger MA, Fabian MA, Biggs WH, Lockhart DJ, Zarrinkar PP, Kuriyan J, Shokat KM. Tuning a three-component reaction for trapping kinase substrate complexes. J Am Chem Soc. 2008; 130:17568–17574.10.1021/ja807066f [PubMed: 19053485] Stöber P, Schelhaas M, Nägele E, Hagenbuch P, Rétey J, Waldmann H. Synthesis of characteristic lipopeptides of the human N-Ras protein and their evaluation as possible inhibitors of protein farnesyl transferase. Bioorg Med Chem. 1997; 5:75–83. [PubMed: 9043659] Tarrant MK, Rho HS, Xie Z, Jiang YL, Gross C, Culhane JC, Yan G, Qian J, Ichikawa Y, Matsuoka T, Zachara N, Etzkorn FA, Hart GW, Jeong JS, Blackshaw S, Zhu H, Cole PA. Regulation of CK2 by phosphorylation and O-GlcNAcylation revealed by semisynthesis. Nat Chem Biol. 2012; 8:262–269.10.1038/nchembio.771 [PubMed: 22267120] Thinon E, Serwa RA, Broncel M, Brannigan JA, Brassat U, Wright MH, Heal WP, Wilkinson AJ, Mann DJ, Tate EW. Global profiling of co- and post-translationally N-myristoylated proteomes in human cells. Nat Commun. 2014; 5:4919.10.1038/ncomms5919 [PubMed: 25255805] Triola G, Brunsveld L, Waldmann H. Racemization-Free Synthesis of S-Alkylated Cysteines via Thiol-ene Reaction. The Journal of Organic Chemistry. 2008; 73:3646–3649.10.1021/jo800198s [PubMed: 18376863] Ubersax JA, Woodbury EL, Quang PN, Paraz M, Blethrow JD, Shah K, Shokat KM, Morgan DO. Targets of the cyclin-dependent kinase Cdk1. Nature. 2003; 425:859–864.10.1038/nature02062 [PubMed: 14574415] Valkevich EM, Guenette RG, Sanchez NA, Chen YC, Ge Y, Strieter ER. Forging Isopeptide Bonds Using Thiol–Ene Chemistry: Site-Specific Coupling of Ubiquitin Molecules for Studying the Activity of Isopeptidases. J Am Chem Soc. 2012; 134:6916–6919.10.1021/ja300500a [PubMed: 22497214] van der Heden van Noort GJ, van der Horst MG, Overkleeft HS, van der Marel GA, Filippov DV. Synthesis of mono-ADP-ribosylated oligopeptides using ribosylated amino acid building blocks. J Am Chem Soc. 2010; 132:5236–5240.10.1021/ja910940q [PubMed: 20232863] van Kasteren SI, Kramer HB, Jensen HH, Campbell SJ, Kirkpatrick J, Oldham NJ, Anthony DC, Davis BG. Expanding the diversity of chemical protein modification allows post-translational mimicry. Nature. 2007; 446:1105–1109.10.1038/nature05757 [PubMed: 17460675] Verdin E, Ott M. 50 years of protein acetylation: from gene regulation to epigenetics, metabolism and beyond. Nat Rev Mol Cell Biol. 2015; 16:258–264.10.1038/nrm3931 [PubMed: 25549891] Vila-Perelló M, Muir TW. Biological Applications of Protein Splicing. Cell. 2010; 143:191– 200.10.1016/j.cell.2010.09.031 [PubMed: 20946979] Virdee S, Kapadnis PB, Elliott T, Lang K, Madrzak J, Nguyen DP, Riechmann L, Chin JW. Traceless and site-specific ubiquitination of recombinant proteins. J Am Chem Soc. 2011; 133:10708– 10711.10.1021/ja202799r [PubMed: 21710965]


Chuh et al.

Page 35


Virdee S, Ye Y, Nguyen DP, Komander D, Chin JW. Engineered diubiquitin synthesis reveals Lys29isopeptide specificity of an OTU deubiquitinase. Nat Chem Biol. 2010; 6:750–757.10.1038/ nchembio.426 [PubMed: 20802491] Vocadlo D, Hang H, Kim E, Hanover J, Bertozzi C. A chemical approach for identifying O-GlcNAcmodified proteins in cells. Proc Natl Acad Sci USA. 2003; 100:9116–9121. [PubMed: 12874386] Wan J, Savas JN, Roth AF, Sanders SS, Singaraja RR, Hayden MR, Yates JR, Davis NG. Tracking brain palmitoylation change: predominance of glial change in a mouse model of Huntington’s disease. 2013; 20:1421–1434.10.1016/j.chembiol.2013.09.018 Wang J, Zhang CJ, Zhang J, He Y, Lee YM, Chen S, Lim TK, Ng S, Shen HM, Lin Q. Mapping sites of aspirin-induced acetylations in live cells by quantitative acid-cleavable activity-based protein profiling (QA-ABPP). Sci Rep. 2015; 5:7896–12.10.1038/srep07896 [PubMed: 25600173] Wang LX. The Amazing Transglycosylation Activity of Endo-β-N-acetylglucosaminidases. Trends Glycosci Glycotechnol. 2011; 23:33–52.10.4052/tigg.23.33 [PubMed: 25309039] Wang LH, Besirli CG, Johnson EM. Mixed-lineage kinases: a target for the prevention of neurodegeneration. Annu Rev Pharmacol Toxicol. 2004; 44:451–474.10.1146/annurev.pharmtox. 44.101802.121840 [PubMed: 14744254] Wang P, Dong S, Shieh JH, Peguero E, Hendrickson R, Moore MAS, Danishefsky SJ. Erythropoietin Derived by Chemical Synthesis. Science. 2013; 342:1357–1360.10.1126/science.1245095 [PubMed: 24337294] Wang R, Islam K, Liu Y, Zheng W, Tang H, Lailler N, Blum G, Deng H, Luo M. Profiling genomewide chromatin methylation with engineered posttranslation apparatus within living cells. J Am Chem Soc. 2013; 135:1048–1056.10.1021/ja309412s [PubMed: 23244065] Wang YS, Wu B, Wang Z, Huang Y, Wan W, Russell WK, Pai PJ, Moe YN, Russell DH, Liu WR. A genetically encoded photocaged Nepsilon-methyl-L-lysine. Mol Biosyst. 2010; 6:1557– 1560.10.1039/c002155e [PubMed: 20711534] Wang Z, Chinoy ZS, Ambre SG, Peng W, McBride R, de Vries RP, Glushka J, Paulson JC, Boons GJ. A General Strategy for the Chemoenzymatic Synthesis of Asymmetrically Branched N-Glycans. Science. 2013; 341:379–383.10.1126/science.1236231 [PubMed: 23888036] Weikart ND, Mootz HD. Generation of site-specific and enzymatically stable conjugates of recombinant proteins with ubiquitin-like modifiers by the Cu(I)-catalyzed azide-alkyne cycloaddition. ChemBioChem. 2010; 11:774–777.10.1002/cbic.200900738 [PubMed: 20209558] Willnow S, Martin M, Lüscher B, Weinhold E. A selenium-based click AdoMet analogue for versatile substrate labeling with wild-type protein methyltransferases. ChemBioChem. 2012; 13:1167– 1173.10.1002/cbic.201100781 [PubMed: 22549896] Wilson JP, Raghavan AS, Yang YY, Charron G, Hang HC. Proteomic analysis of fatty-acylated proteins in mammalian cells with chemical reporters reveals S-acylation of histone H3 variants. Mol Cell Proteomics. 2011; 10:M110.001198.10.1074/mcp.M110.001198 [PubMed: 21076176] Winder WW, Hardie DG. AMP-activated protein kinase, a metabolic master switch: possible roles in type 2 diabetes. Am J Physiol. 1999; 277:E1–10. [PubMed: 10409121] Wolfert MA, Boons GJ. adaptive immune activation: glycosylation doesmatter. Nat Chem Biol. 2013; 9:776–784.10.1038/nchembio.1403 [PubMed: 24231619] Woo CM, Iavarone AT, Spiciarich DR, Palaniappan KK, Bertozzi CR. Isotope-targeted glycoproteomics (IsoTaG): a mass-independent platform for intact N- and O-glycopeptide discovery and analysis. Nat Meth. 201510.1038/nmeth.3366 Wright MH, Clough B, Rackham MD, Rangachari K, Brannigan JA, Grainger M, Moss DK, Bottrill AR, Heal WP, Broncel M, Serwa RA, Brady D, Mann DJ, Leatherbarrow RJ, Tewari R, Wilkinson AJ, Holder AA, Tate EW. Validation of N-myristoyltransferase as an antimalarial drug target using an integrated chemical biology approach. Nature Chemistry. 2014; 6:112– 121.10.1038/nchem.1830 Wright TH, Brooks AES, Didsbury AJ, Williams GM, Harris PWR, Dunbar PR, Brimble MA. Direct Peptide Lipidation through Thiol-Ene Coupling Enables Rapid Synthesis and Evaluation of SelfAdjuvanting Vaccine Candidates. Angew Chem Int Ed. 2013; 52:10616–10619.10.1002/anie. 201305620


Chuh et al.

Page 36


Xie J, Supekova L, Schultz PG. A genetically encoded metabolically stable analogue of phosphotyrosine in Escherichia coli. ACS Chem Biol. 2007; 2:474–478.10.1021/cb700083w [PubMed: 17622177] Yaffe MB. Phosphotyrosine-binding domains in signal transduction. Nature Publishing Group. 2002; 3:177–186.10.1038/nrm759 Yaffe MB, Elia AE. Phosphoserine/threonine-binding domains. Curr Opin Cell Biol. 2001; 13:131– 138. [PubMed: 11248545] Yang YY, Ascano JM, Hang HC. Bioorthogonal Chemical Reporters for Monitoring Protein Acetylation. J Am Chem Soc. 2010; 132:3640–3641.10.1021/ja908871t [PubMed: 20192265] Yang YY, Grammel M, Hang HC. Identification of lysine acetyltransferase p300 substrates using 4pentynoyl-coenzyme A and bioorthogonal proteomics. Bioorg Med Chem Lett. 2011; 21:4976– 4979.10.1016/j.bmcl.2011.05.060 [PubMed: 21669532] Yin L, Krantz B, Russell NS, Deshpande S, Wilkinson KD. Nonhydrolyzable Diubiquitin Analogues Are Inhibitors of Ubiquitin Conjugation and Deconjugation †. Biochemistry. 2000; 39:10001– 10010.10.1021/bi0007019 [PubMed: 10933821] Yount JS, Moltedo B, Yang YY, Charron G, Moran TM, López CB, Hang HC. Palmitoylome profiling reveals S-palmitoylation–dependent antiviral activity of IFITM3. Nat Chem Biol. 2010; 6:610– 614.10.1038/nchembio.405 [PubMed: 20601941] Zaro BW, Yang YY, Hang HC, Pratt MR. Chemical reporters for fluorescent detection and identification of O-GlcNAc-modified proteins reveal glycosylation of the ubiquitin ligase NEDD4-1. Proc Natl Acad Sci USA. 2011; 108:8146–8151.10.1073/pnas.1102458108 [PubMed: 21540332] Zhang C, Kenski D, Paulson J, Bonshtien A, Sessa G, Cross J, Templeton D, Shokat K. A second-site suppressor strategy for chemical genetic analysis of diverse protein kinases. Nat Meth. 2005; 2:435–441. Zhang FL, Casey PJ. Protein prenylation: molecular mechanisms and functional consequences. Annu Rev Biochem. 1996; 65:241–269.10.1146/annurev.bi.65.070196.001325 [PubMed: 8811180] Zhang J, Yang PL, Gray NS. Targeting cancer with small molecule kinase inhibitors. Nat Rev Cancer. 2009; 9:28–39.10.1038/nrc2559 [PubMed: 19104514] Zhang MM, Wu PYJ, Kelly FD, Nurse P, Hang HC. Quantitative control of protein S-palmitoylation regulates meiotic entry in fission yeast. PLoS Biol. 2013; 11:e1001597.10.1371/journal.pbio. 1001597 [PubMed: 23843742] Zhang Z, Shen K, Lu W, Cole PA. The role of C-terminal tyrosine phosphorylation in the regulation of SHP-1 explored via expressed protein ligation. J Biol Chem. 2003; 278:4668–4674.10.1074/ jbc.M210028200 [PubMed: 12468540] Zheng B, DeRan M, Li X, Liao X, Fukata M, Wu X. 2-Bromopalmitate Analogues as Activity-Based Probes To Explore Palmitoyl Acyltransferases. J Am Chem Soc. 2013; 135:7082–7085.10.1021/ ja311416v [PubMed: 23631516] Zheng B, Zhu S, Wu X. Clickable Analogue of Cerulenin as Chemical Probe to Explore Protein Palmitoylation. ACS Chem Biol. 2015; 10:115–121.10.1021/cb500758s [PubMed: 25322207] Zhu Y, van der Donk WA. Convergent Synthesis of Peptide Conjugates Using Dehydroalanines for Chemoselective Ligations. Org Lett. 2001; 3:1189–1192.10.1021/ol015648a [PubMed: 11348191]

Author Manuscript Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 37

Author Manuscript Author Manuscript Figure 1. Encoding and decoding posttranslational modifications (PTMs)

Author Manuscript

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.


Chuh et al.

Page 38


Figure 2. Encoding protein phosphorylation

(A) A family of ~500 kinases will transfer a phosphate group to certain amino acid sidechains, 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


Chuh et al.

Page 39

Author Manuscript

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.

Author Manuscript Author Manuscript Author Manuscript Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 40


Figure 3. Decoding protein phosphorylation

Author Manuscript

(A) Development of analog-sensitive kinases using a bump-hole strategy. Wild-type kinases are incapable of using the “bumped” ATP analog N6-benzyl 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-benzylATPɣS, 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.


Chuh et al.

Page 41


Figure 4. Protein glycosylation

Author Manuscript

(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 synthesized after peptide synthesis through the coupling of an aspartic acid residue to a glycosyl-amine or amine equivalent. (C) Enzymatic installation of large Nlinked 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.


Chuh et al.

Page 42

Author Manuscript 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 δ-mercapto-lysine residue. The δ-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 δ-thiol group is then removed by chemical desulfurization. (D) A δ-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.


Chuh et al.

Page 43


Figure 6. Lipidation

(A) Proteins can be modified by several types of lipids, including myristoylation at the Ntermini 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 Nethylmaleimide. Any palmitate thioesters are then cleaved using hydroxylamine and the resulting free thiols can be reacted with a variety of electrophilic tags.

Author Manuscript Author Manuscript Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 44


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.

Author Manuscript Author Manuscript Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chuh et al.

Page 45


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


Chuh et al.

Page 46

Author Manuscript

to a variety of protein side chains and then subsequently polymerized to form long polyADP-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 ADPribosylation reporters for use in cell lysates.

Author Manuscript Author Manuscript Author Manuscript Cell Chem Biol. Author manuscript; available in PMC 2017 January 21.

Chemical methods for decoding cytosine modifications in DNA.

Chemical proteomics approaches to examine novel histone posttranslational modifications.

Posttranslational Modifications and the Immunogenicity of Biotherapeutics.

Protein synthesis, posttranslational modifications, and aging.

Novel encoding methods for DNA-templated chemical libraries.

Dynamics of posttranslational modifications of p53.

Mutant TP53 posttranslational modifications: challenges and opportunities.

The Functional Significance of Posttranslational Modifications on Polo-Like Kinase 1 Revealed by Chemical Genetic Complementation.

Posttranslational modifications of histone deacetylases: implications for cardiovascular diseases.

Autoantibodies to posttranslational modifications in rheumatoid arthritis.

Chromogranin A: posttranslational modifications in secretory granules.

Encoding and decoding nonverbal cues of emotion.

Causal interpretation rules for encoding and decoding models in neuroimaging.

Data-driven HRF estimation for encoding and decoding models.

Immunogenicity of Biotherapeutics: Causes and Association with Posttranslational Modifications.

Targeted proteomics of myofilament phosphorylation and other protein posttranslational modifications.

Posttranslational modifications of proopiomelanocortin in vertebrates and their biological significance.

Visualizing posttranslational and epigenetic modifications of endogenous proteins in vivo.

Encoding and decoding spatio-temporal information for super-resolution microscopy.

Decoding liver injury: A regulatory role for histone modifications.

Posttranslational Modifications and Clearing of α-Synuclein Aggregates in Yeast.

Chemical shift encoding-based water-fat separation methods.

Chemical biology strategies for posttranslational control of protein function.

Posttranslational modifications of α-tubulin in alzheimer disease.