Molecular Perspectives on Protein Adenylylation - ACS Chemical

Reviews pubs.acs.org/acschemicalbiology

Molecular Perspectives on Protein Adenylylation Christian Hedberg*,†,‡ and Aymelt Itzen*,§ †

Chemical Biology Center (KBC), Institute of Chemistry, Umeå University, Umeå, 90187, Sweden Max Planck Institute of Molecular Physiology, Department of Chemical Biology, Dortmund 44227, Germany § Center for Integrated Protein Science Munich, Chemistry Department, Technische Universität München, Lichtenbergstr. 4, 85748 Garching, Germany ‡

ABSTRACT: In the cell, proteins are frequently modified covalently at specific amino acids with post-translational modifications, leading to a diversification of protein functions and activities. Since the introduction of high-resolution mass spectrometry, new post-translational modifications are constantly being discovered. One particular modification is the adenylylation of mammalian proteins. In adenylylation, adenosine triphosphate (ATP) is utilized to attach an adenosine monophosphate at protein threonine or tyrosine residues via a phosphodiester linkage. Adenylylation is particularly interesting in the context of infections by bacterial pathogens during which mammalian proteins are manipulated through AMP attachment via secreted bacterial factors. In this review, we summarize the role and regulation of enzymatic adenylylation and the mechanisms of catalysis. We also refer to recent methods for the detection of adenylylated proteins by modification-specific antibodies, ATP analogues equipped with chemical handles, and mass spectrometry approaches. Additionally, we review screening approaches for inhibiting adenylylation and briefly discuss related modifications such as phosphocholination and phosphorylation.

T

be attacked, resulting in adenylylation and pyrophosphorylation, respectively. In this review, we will focus on protein adenylylation (Figure 1A). Protein adenylylation is a PTM that is simultaneously old and new. Earl Stadman discovered the reversible adenylylation of bacterial glutamine synthetase (GS) by GS adenylyltransferase (GS ATase) as part of the metabolic regulation of glutamine synthesis (reviewed in ref 1). For many years, this discovery has been the only case of protein adenylylation in the literature; however, the field was revived when several groups (Orth, Dixon, Itzen, and Goody) observed the adenylylation (also referred to as AMPylation) of mammalian Rho and Rab proteins in response to bacterial infections about 40 years later.2−4 These contributions have received considerable attention since they suggested that adenylylation might be involved in the establishment of various bacterial infections. We will therefore summarize the currently known mechanisms and the potential regulation of protein adenylylation, comment on the reversibility of this PTM, and discuss approaches and challenges in the detection of modified protein substrates. However, we will not allude to the adenylylation reactions that are part of the activation of protein/amino acid carboxylates

he functional repertoire of proteins is greatly diversified by post-translational modifications (PTMs). Just like a smartphone that can be adjusted in its functionality/ applicability by installation of individual software components (i.e., apps), a protein can and/or must be equipped with covalently attached functional groups to finally reach its predetermined physiologically relevant form after translation into the premature state. This covalent addition of groups is referred to as PTM. Although PTMs such as proteolysis and phosphorylation have been known for a long time, the full scope of the biological repertoire of PTMs has only been possible to address in an efficient way since the introduction of high resolution mass spectrometry (MS) methods for tryptic fragments of proteins. Since then, the list of PTMs and the modified amino acids is ever growing. Many PTMs are brought about reversibly (or irreversibly) by protein modifying enzymes that utilize a metabolite with high group transfer potential. Ribonucleotides are particularly privileged molecules in this respect, as they are abundant and contain four high-energy bonds with electrophilic centers: two phosphoanhydrides, a phosphoester, and a glycosidic linkage. The phosphoanhydride bonds are far more reactive than the others and make up for most of the relevant transfer reactions. They are amenable to attack from nucleophilic groups of amino acid side chains and the amino or carboxy terminal parts of proteins by enzyme catalysis. The most prominent nucleotidebased PTM is phosphorylation, which results from a nucleophilic attack of a protein at the γ-phosphate of adenosine triphosphate (ATP). Also the α- and β-phosphate positions can © XXXX American Chemical Society

Special Issue: Post-Translational Modifications Received: October 21, 2014 Accepted: December 8, 2014

A

dx.doi.org/10.1021/cb500854e | ACS Chem. Biol. XXXX, XXX, XXX−XXX

ACS Chemical Biology

Reviews

Figure 1. Mechanisms of adenylylation. (A) Reaction scheme of protein adenylylation for threonine and tyrosine side chains. The adenosine is transferred along with the α-phosphate to the amino acid side chain, forming the new phosphodiester bond. Pyrophosphate is a byproduct of the reaction and is subsequently hydrolyzed by pyrophosphatase yielding two phosphates. (B) Mechanism of adenylylation by DNA-polymerase-β-like enzymes, exemplified by the proposed mechanism of DrrA.9 Three acidic side chains are involved in Mg2+ coordination; a fourth one acts as general base to deprotonate the incoming nucleophilic side chain of the protein substrate (e.g., Y77 of Rab1b). (C) Mechanism of adenylylation by Ficdomains inferred from published crystal structures.13,14,16 The catalytic histidine may coordinate the 5′ prime oxygen and act as a general base for deprotonation of the protein substrate (e.g., Thr35 of Rac1 modified by VopS). The figure underlines the significance of the Fic-motif in coordinating the phosphates of the nucleotide (green, ATP; Mg2+, magnesium ion; red arrows, reaction step during adenylylation).

following β sheet. Although this family consists of several members, only the enzymes DrrA from Legionella pneumophila and the GS ATase from bacteria have been reported to act in protein adenylylation.2,6,8 The catalytic mechanism of AMP transfer has recently been elucidated in detail and suggests that the second aspartate (D112DrrA) of the DrrA motif G−X11− [ED]−X−[ED] acts as a general base to deprotonate the target tyrosine Y77 of Rab1b and thereby prepares it for nucleophilic attack on the α-phosphate of ATP.9 Interestingly, DrrA also accepts alternative nucleotides, such as CTP and GTP, albeit with lower catalytic activities (60% and 14%, respectively).10 In contrast to the narrow family of DNA-polymerase-β-like adenylyl transferases, the Fic (filamentation induced by cAMP) family of enzymes currently comprises roughly 3000 structurally and functionally related members.11 The unifying structural element of this family is a characteristic helical bundle of six αhelices that surround a loop region with a highly conversed amino acid sequence motif HXFX(D/E)(A/G)N(G/K)R (referred to as the Fic motif), the presence of which is sufficient to identify a Fic protein by sequence analysis (Figure 1C).11,12 Often, the basic core is surrounded by structural appendages that assist in protein substrate binding.12−15 The catalytic loop serves two functions: On the one hand, the conserved histidine acts as general base to deprotonate a

(e.g., ubiquitination, protein/peptide biosynthesis) or of the polymerization of DNA or RNA (see ref 5 for information).

■

ENZYMATIC PROTEIN ADENYLYLATION In the process of enzymatic adenylylation, ATP is utilized to covalently transfer an AMP moiety to an amino acid side chain (Figure 1A). Currently, tyrosine and threonine are known targets,2−4 but also serine could, in principle, be conceivable as a stable modification. This is a highly exergonic reaction since ATP contains two anhydride linkages that are converted into a phosphodiester product and a pyrophosphate. Under physiological conditions, the pyrophosphate is subsequently hydrolyzed enzymatically by pyrophosphatase, thus shifting the equilibrium completely to the product side. Adenylylating enzymes are currently grouped into two different families according to their structural properties and enzymatic mechanism: the DNA-β-polymerase-like and the Fic family of adenylylating enzymes. The DNA-polymerase-β-like family of nucleotidyl transferases contains a structurally characteristic three-stranded β-sheet involved in magnesium ion coordination and phosphate binding (Figure 1B).6−8 In addition, it also contains the degenerate sequence motif G− X11−[ED]−X−[ED] in which the aspartates coordinate two Mg2+’s together with another aspartate or glutamate from the B



Reviews

pneumophila adenylylates switch II Y77 of Rab1b,2 VopS of Vibrio parahaemolyticus modifies switch I T35 of Rac,3 and IbpA of Histophilus somni attaches AMP to switch I Y34 of RhoA (Y32 in Rac and Cdc42).4 Since the switch regions are intimately involved in binding guanosine diphosphate (GDP) or guanosine triphosphate (GTP) and in catalyzing GTPhydrolysis, adenylylation of these loops could hypothetically affect GDP and/or GTP affinities and the rates of GTP conversion. Conclusive data on the consequences of adenylylation on the intrinsic GTPase parameters only exist for Rab1b, where no effects on GTP-hydrolysis or nucleotide binding have been detected.10 However, since adenylylations of Rac1 and RhoA occur on conserved amino acids (Rac1 Y32 vs Rac1 T35) in a different region (switch I), the consequences remain to be tested. The effects of GTPase adenylylation have been analyzed in more detail on the level of protein−protein interactions (Figure 2A). Adenylylation of Rab1b blocks binding to most mammalian and some Legionella proteins but intriguingly maintains the interaction with the Legionella protein LidA and the guanine nucleotide exchange factor (GEF) domain of DrrA.2,10,24−26 Since Rab1b adenylylation profoundly affects the binding of GTPase activating proteins (GAPs) that act in deactivating Rab1b by stimulating GTP-to-GDP conversion, adenylylation has been considered to lock Rab1b in the active, GTP-bound state. Additionally, the block of binding to the recycling factor RabGDI (Rab GDP dissociation inhibitor) suggests a stimulation of membrane binding and thus stabilizes recruitment of Rab1b. 26,27 In the case of RhoA/Rac adenylylation by VopS and IbpA, the analyses of molecular consequences are less detailed. It is unambiguous, however, that adenylylation of these GTPases blocks the interaction with downstream binding partners and thereby inhibits the formation of RhoA-controlled cytoskeletal structures.3,4 It has also been observed recently that adenylylation of Rho family GTPases can affect signaling pathways controlled by NFkB, ERK, and Jnk; inhibit Rho GTPase degradation; and attenuate antimicrobial superoxide production.28 In addition to modifications of mammalian proteins by bacterial Fic enzymes, it has also been demonstrated that the eukaryotic protein FicD (human name: HYPE, hFic) can adenylylate eukaryotic target substrates.4,15,17,29 Thus, adenylylation is not limited to the interaction of bacteria with eukaryotic cells during infections. Experiments with purified proteins and cell lysates demonstrated that HYPE is capable of adenylylating small GTPases of the Rho family at Y34 similar to IbpA.4 Very recently, it has additionally been shown that the HYPE-homologue FicD from Drosophila melanogaster (dFic) causes reversible site specific adenylylation of the endoplasmic reticulum chaperone BiP at the conserved amino acid residue T366 in vivo.29 The enzyme dFic is localized in the endoplasmic reticulum and apparently is involved in fly development.30 Very interestingly, the site of adenylylation at T366 is localized in close proximity to the ATP-binding site of BiP. Since the ATP binding and ATPase activities are important for the function of BiP as a molecular chaperone in the ER,31 adenylylation at T366 may regulate the activity of BiP. It has been observed that BiP adenylylation is decreased during ER stress and correlates with the inactive state of the chaperone.29 However, the exact molecular consequences of BiP adenylylation remain to be determined. The recent crystal structure of human HYPE may provide a basis for further exploring the molecular processes involved in BiP adenylylation.15

specific nucleophilic side chain (e.g., serine, tyrosine) and is thus essential for enzymatic activity.3,4,13,14 On the other hand, the loop properly positions and activates the phosphates of the nucleotide for nucleophilic attack by the protein substrates’ side chain (Figure 1C). Here, the residues (A/G)N(G/K) form an anion acceptor site that may stabilize the developing negative charge on the bound phosphate (α-phosphate in the case of adenylylation by ATP) during nucleophilic attack by the protein substrate.12,13,16 Additionally, the Fic-loops’ positively charged arginines supports this stabilization by coordinating the neighboring phosphate moiety (β-phosphate in the case of adenylylation by ATP). Since the Fic motif is binding the phosphate functionalities, but does not, or only weakly, contribute to the ribose or the nucleobase recognition, the relative orientation of the nucleotide needs to be determined by other structural elements of the enzyme. It has been observed recently that also other nucleotides (e.g., CTP, CDP-choline17,18), and alternative orientations of ATP in the binding pocket are possible, thus resulting in different modifications such as phosphocholination or phosphorylation (see section Related Modifications for further explanations).12,18−21 Currently, the enzymatically catalyzed attachment of nucleotide monophosphates (NMPs) is centered on protein adenylylation. It is, however, conceivable that also the other NMPs (e.g., CMP, etc.) can react with amino acid side chains of proteins. The nucleotidylation of protein targets with cytidine monophosphate (CMP) has been demonstrated previously in vitro with purified enzymes and pure nucleotide triphosphates (NTPs).10,17 Nevertheless, ATP is under cellular conditions the most abundant among the NTPs and therefore likely competes efficiently with the others. It has also been shown in one instance that ATP is the preferred substrate of the adenylylating enzyme DrrA, and consequently only adenylylation has been observed.10 Since nucleotidylation appears to sterically interfere with complex formations negatively (see section Consequences of Adenylylation), the identity of the nucleobase is likely not relevant for the biological effect of the modification. If, however, nucleotidylations could also be involved in generating new sites for intra- or intermolecular interactions such as in phosphorylations, the base may play a decisive role for the physiological output. Nonetheless, this hypothesis remains to be shown.

■

CONSEQUENCES OF ADENYLYLATION Adenylylation of proteins introduces a sterically demanding AMP moiety into proteins and changes the net charge of the target molecule by −1. Consequently, it is expected that this modification alters either intrinsic properties (e.g., stability, enzymatic activity, cofactor binding, etc.) of the substrate or changes its protein interaction profile. The most detailed characterizations of protein adenylylation have been performed on mammalian small GTPases and on the bacterial glutamine synthetase. Bacterial pathogens often secrete enzymes that post-translationally modify small GTPases of mammalian cells due to their pivotal role in controlling and combating infections22 (for a review on small GTPases, see ref 23). Small GTPases contain two highly important regulatory loop regionsreferred to as switch I and switch IIthat mediate the interaction with upstream regulators and downstream effectors. In three instances, adenylylation of GTPases has been observed to occur in these switch regions: The enzyme DrrA from L. C



Reviews

enzyme by altering catalytic activities: The activity of bacterial glutamine synthetase (GS)a dodecameric enzyme that generates glutamine from glutamate and ammoniais regulated by reversible adenylylation.1,32,33 Each of the 12 GS subunits can be adenylylated by the enzyme GS ATase at Y398GS leading to gradual inhibition of glutamine synthesis. However, this is the only reported example of direct enzymatic activity changes due to adenylylation. One could also speculate about further consequences of protein adenylylation. In analogy to O-GlcNAcylation,34 adenylylation may block sites of protein phosphorylation reversibly or irreversibly. However, none of the currently identified adenylylation sites in Rab1, RhoA, GS ATase, or BiP have been demonstrated to be subject to phosphorylations, yet. VopS adenylylates the protein LyGDI in a region that can be also phosphorylated by Src kinase.35 Adenylylation interfered negatively with phosphorylation even though the sites of adenylylation and phosphorylation are not identical. A similar masking effect with respect to phosphorylation sites has been observed with the plant kinases RipK and Bik1 that are uridylylated (i.e., modified with uridine monophosphate) by AvrAC.36 These findings demonstrate that there is crosstalk between adenylylation and other PTMs.

■

REGULATION OF ADENYLYLATION Since protein adenylylation has dramatic consequences for enzymatic activity (e.g., GS ATase) or protein interaction profiles (e.g., DrrA, VopS, IbpA), it is conceivable that the activity of adenylylating enzymes or the level of adenylylation is controlled by internal or external parameters. Interestingly, it has been found that the catalytic activity of a group of Fic enzymes is regulated through the presence of an inhibitory α-helix (αinh):15,16,37 This helix contains a (S/ T)XXXE(G/N) sequence motif in which the conserved glutamate protrudes into the phosphate binding pocket of catalytic site of Fic enzymes and thereby sterically and electrostatically obstructs ATP binding and phosphate positioning (Figure 2B, top panel).15,16 The inhibitory helix can be part of an antitoxin module (class I αinh) provided in trans or can be present at the N-terminus (class II αinh) or the C-terminus (class III αinh) of the Fic enzyme. However, the mechanism of the release of inhibition is currently unknown. The presence of the inhibitory helix nevertheless suggests that such Fic enzymes require secondary factors acting in the installation or release of the inhibition. Additionally, the recent structural and biochemical characterization of the human Fic proteins HYPE suggests that the adenylylation activity may also be regulated through enzyme dimerization for some family members:15 One amino acid substitution interfering with homo dimerization decreased the autoadenylylation activity of HYPE. A different approach to control protein adenylylation is employed by infection with L. pneumophila. During infection, Legionella bacteria secrete about 300 different proteins directly from the cytosol of the bacterium to the cytosol of the host cell.38 The adenylylating activity of Legionella DrrA is counteracted by the deadenylylating bacterial enzyme SidD.39−41 SidD hydrolytically cleaves the AMP from Rab1b and thereby restores the unmodified proteins (Figure 2B, middle panel). In order to prevent futile cycles of adenylylation and deadenylylation, DrrA (0−4 h post infection) and SidD (>1 h post infection) appear to be secreted at different time points in the course of infection.42 Thus, immediately after infection Rab1b is adenylylated by DrrA and maintained in this

Figure 2. Consequences and regulation of protein adenylylation. (A) Adenylylation of small GTPases (Rho, Rab) negatively interferes with binding to regulatory factors (e.g., GAPs, GDI) and effectors. Interestingly, binding to the bacterial Legionella protein LidA is still possible for adenylylated Rab1. (B) Levels of adenylylation regulation. Top panel (inhibitory regulation): Selected Fic enzymes are regulated through an inhibitory α-helix (αinh) that sterically and electrostatically interferes with ATPs’ phosphate binding via a conserved acidic side chain. Middle panel (reversible adenylylation): Protein adenylylation can be regulated through reversion of the modification. The Legionella enzyme SidD hydrolytically restores unmodified Rab1 during bacterial infection with AMP as a byproduct. In contrast, the deadenylyating activity at the N-terminus of GS ATase operates via phosphorolysis of modified glutamine synthetase (GS), leading to the production of unmodified GS and ADP. Bottom panel (enzyme activity control): The enzymatic activity of GS ATase is controlled by the regulatory protein PII (PII). PII binds to a regulatory site (R) that connects the Nterminal AMP-removase and the C-terminal AMP-transferase domains of GS ATase. If PII is post-translationally modified with UMP, it stimulates the removase and inhibits the transferase activities of GS ATase. Unmodified PII, however, leads to the opposite effects.

Initially, protein adenylylation was reported by Earl C. Stadman to function in enzymatic control of a metabolic D



Reviews

Figure 3. Chemical methods for the analysis of adenylylation. (A) Designing building blocks with 2′,3′ isopropylidene protection of the ribofuranose moiety allows for efficient solid-phase peptide synthesis of adenylylated peptides relying on global acidolytic deprotection/resin cleavage. In case of Ser/Thr, the phosphodiester linkage is left unprotected for the synthesis sequence to prevent ß-elimination. (B) For metabolic labeling of Ficdomain substrates, different functionalized ATP analogues have been employed. N6-propargyl ATP (left) has been developed for pulldown strategies employing biotin-azide click reaction. Fluorescein-ATP on the other hand (right) allows for the direct detection of adenylylated proteins by in-gel fluorescence. (C) Fragmentation patterns for adenylylated amino acids under Ms/Ms conditions. Adenylylated tyrosine (left) gives rise to distinctly different reporter ions compared to serine and threonine (right). (D) Inhibitors of VopS adenylylation. By using Fluorescein-ATP in a fluorescence polarization assay, it was possible to identify calmidazolium (right) and other derivatives (left) possessing the parent scaffold as VopS inhibitors. E



Reviews

state.42 At about 3−4 h post infection, DrrA expression would cease, whereas SidD levels increase gradually, leading to effective global Rab1b deadenylylation.39 Therefore, reversible adenylylation at least in this instance is regulated through the expression, secretion, and availability of the modifying and demodifying enzymes. Adenylylations of RhoA proteins by VopS or IbpA have so far not been observed to be reversible under physiological conditions, but Rac1 adenylylated on Y32Rac1 by IbpA can be hydrolytically cleaved by snake venom phosphodiesterase I in vitro.4 The regulation of reversible adenylylation via counteracting enzymatic mechanisms is also realized in the control of the bacterial GS ATase (Figure 2B, middle panel). GS ATase is composed of three functionally distinct domains: an N-terminal deadenylylation domain (also referred to as adenylyl removase, AR), a central regulatory site (R), and a C-terminal adenylyl transferase (ATase) domain. 43−46 The switch between adenylylation (ATase) and deadenylylation (AR) is made by a regulatory factor referred to as PII that can bind to the R site (Figure 2 B, bottom panel).6 Interestingly, PII is also regulated by PTMs that determine the activity state of the protein. If PII is uridylylated (i.e., modified with uridine monophosphate) binding to the R site causes the AR activity of GS ATase to dominate, leading to GS deadenylylation.47,48 If, however, PII is not uridylylated, the binding to the R domain stimulates the adenylylation activity of GS ATase.47,48 Thus, elaborate cycles of reversible uridylylation and adenylylation regulate the activity of GS. In contrast to the hydrolytic deadenylylation reaction described for Legionella SidD,39,40 deadenylylation of GS occurs by AR-catalyzed phosphorolysis yielding adenosine diphosphate (ADP) (not AMP) as a byproduct.49 In summary, there are many ways to regulate protein adenylylation, but there is no general picture emerging from previous examples.

sequences employing more forcing coupling conditions remains to be elucidated. In conclusion, 2′,3′-isopropylidene protection of the adenosine moiety, together with solubility-enhancing bisboc protection at N6 provides the best flexibility for synthesis of adenylylated peptides (Figure 3A). Detection by Chemical Handles. Chemical handles (e.g., fluorophores, affinity tags) placed at strategically convenient groups of the utilized substrate compounds can dramatically simplify the detection of post-translationally modified proteins (for reviews, see refs 53 and 54). In particular, the use of alkyne or azide groups that can be subsequently reacted using bioorthogonal copper(I)-catalyzed azide−alkyne cycloaddition (CuAAC or click chemistry) has revolutionized the detection of PTMs. This concept has been realized previously for the fluorescence imaging and enrichment of nascent mRNA transcript through metabolic labeling with alkyne-bearing N6propargyl adenosine that could be utilized by mammalian RNA polymerases. 55Recently, an alkynyl tag (propargyl) has been introduced at the N6 position of the adenine of ATP (N6pATP) and combined with the click reaction to also identify adenylylated proteins.56 Similarly, ATP derivatives with a fluorophore at the adenine N6 NH2 could be utilized to successfully label VopS substrates and to detect previously unrecognized modified proteins.57 Even though this strategy is minimally invasive, it may not be generally applicable due to the potential involvement of the N6 position in binding to the enzymatic pocket. Thorough investigation of adenylylation by Legionella DrrA and structural investigations of NmFic from Neisseria meningitis and IbpA from H. somni suggested that this indeed may be the case.10,14,16 It also has been proposed that the N6 position may be crucial for some adenylylating enzymes (DrrA, IbpA),10,17 whereas other Fics are rather unselective for their nucleotide (VopS). Consequently, dramatically different Fic concentrations have to be applied in order to achieve similar turnover rates.56 A systematic analysis of other positions at the nucleotide will thus be required to identify a more generic nucleotide substrate (Figure 3B). N6pATP has been used successfully for profiling the activities and specificities of selected Fic proteins despite its limitations for serving as a generic ATP surrogate. In a conceptionally very appealing study, it was possible to profile the adenylylating activity of VopS and IbpA against a library of expressed proteins using N6pATP.35 A human cDNA library has been expressed on a nucleic acid programmable protein array (NAPPA) in a microarray format using cell-free transcription/translation. The cDNA has been spotted with spatial control, and the expressed GST or Flag fusion proteins were captured in the vicinity of the cDNA via the binding to immobilized GST or Flag specific antibodies, respectively. More than 20 different previously unrecognized substrates could subsequently be identified after incubation with the adenylylating enzymes in the presence of N6pATP, followed by subsequent click reaction with azido-rhodamine.35 The advantages over antibody based detection methods are lower levels of background noise resulting from unspecific binding and a higher sensitivity. However, even if a generically applicable and potentially clickable ATP analogue can be derived for enzymatic adenylylations, its usage is likely restricted to in vitro analyses. A profound limitation of nucleotide analogues is their insignificant uptake into live cells due to the high negative charge of the phosphates that prevent membrane crossing. Additionally, the adenosine (e.g., N6-propargyl adenosine) or

■

DETECTION OF ADENYLYLATION Synthesis of Adenylylated Peptides. For fundamental investigation of protein adenylylation, synthetic reference material is required. The MS method development and generation of specific AMP antibodies is dependent on efficient synthesis methods to prepare pure peptides bearing adenylylated amino acid side chains (for a historic approach on the synthesis of adenylylated peptides, see ref 5). Recent approaches have solved the problems of poor synthesis yields, leading to high quality peptides.50 The synthesis strategy used 2′,3′-isopropylidene acetal protective groups of adenosine in combination with deactivating bis-boc protection at the adenine nitrogen. The classical O-cyanoethyl protection group was used for stabilization of the phosphodiester linkage followed by in situ deprotection during the first Fmoc removal, thus stabilizing the phosphodiester linkage in the monoanionic form. Standard Fmoc-solid phase peptide synthesis was subsequently used with the tyrosine building block for the generation of adenylylated peptides. Adenylylated serine and threonine residues have also been generated using a strategy based on the unprotected phosphodiesters with the allyl protective group of the phosphodiester linkage being cleaved at the end of the building block synthesis.51 A similar building block strategy relied on the unprotected N6 nitrogen of the adenine. Due to the low nucleophilicity of the adenine NH2 relative to the α-amino group of amino acids, the lack of protection is claimed to be compatible with standard peptide synthesis methods.52 Whether this is valid for longer and more complex peptide F



Reviews

fragmentation of the adenylylated peptides, the important characteristic ions of adenine (m/z 136) and adenosine (m/z 250) or AMP (m/z 348) are not visible and can therefore not be used to positively identify adenylylated peptides. However, due to the less noisy spectra, it appears that search engines can more easily identify the correct peptides with this method, although the results need to be reviewed critically (Figure 3C). Small Molecule Inhibition of Adenylyl Transferases. It has recently been suggested that blocking adenylylation may be an effective way to combat infections; however, no small molecule inhibitors have been available for any of the adenylyl transferases. Thompson and co-workers recently developed a fluorescence-polarization-based high-throughput screening assay and used it to discover the first inhibitors of protein adenylylation.66 Validation in a secondary biochemical screen was performed by Western blotting with α-AMP-threonine antibody.60,66 They identified the first small molecule inhibiting VopS (e.g., calmidazolium, GW7647, and MK886) with inhibitory constants (Ki) ranging from 6 to 50 μM and an at least 30-fold selectivity versus HYPE,4 the only known human (putative) adenylyl transferase. However, cellular toxicity of the identified compounds proved to be high. More potent inhibitors must be designed before proof-of-concept can be achieved in an infection system (Figure 3D).

adenosine phosphate (e.g., N6pATP) derivatives compete against the high endogenous (millimolar) concentrations of ATP in the cell or cell lysates, and thus only minor modification levelsif at allcan be expected. Antibody based methods may hold a potential solution to these shortcomings. Antibody-based Methods. The high selectivity and substrate affinity of antibodies makes them invaluable tools for the analysis of PTMs. Antibodies may be used for the detection and isolation of adenylylated proteins from cells and cell lysates. The first polyclonal α-AMP antibodies were generated more than three decades ago.58,59 Only recently, however, high quality α-AMP antibodies could be obtained by using adenylylated peptides for immunization.50,60 Modified peptides are ideally suited over modified proteins for immunization since they prevent the parallel production of antibodies that are directed against undesired protein epitopes. Additionally, adenylylation significantly alters protein properties given the polar character of AMP (anionic phosphodiester), its hydrophobicity (nucleobase), and the size of the modification. Thereby, raising AMP antibodies against a selected amino acid instead of a peptide sequence should be facilitated. The availability of amino acid building blocks for incorporation of both adenylylated Ser, Thr, and Tyr residues in peptides have simplified the production of specific antibodies.50,51 In this respect, experiments for enrichment of PTM-modified proteins from cell lysates could be performed after general proteolytic digest of the cell lysates to both increase the accessibility of the PTMs and decrease nonspecific binding of the antibodies to random proteins in the lysate.61 The recently derived antibodies have high affinity to Tyradenylylated proteins, with weak Thr-AMP-binding.50 However, Thr-AMP antibodies raised in a similar way did only weakly recognize adenylylated Tyr residues but detected Thr adenylylation on a protein level.60 Mass Spectrometry Techniques. Detection of adenylylated peptides have been reported previously using MS in different fragmentation modes. The peptide antibiotics lincomycin and clindamycin could be inactivated by the addition of AMP and have been investigated using MS.62,63 In a previous MS study, synthetic nontryptic adenylylated peptides were utilized to reveal several new fragments and neutral losses.64 The fragmentation of adenylylated tryptic peptides derived from adenylylated proteins has also recently been investigated:65 Adenylylated peptides disintegrate at different parts of the AMP in response to distinctive fragmentation techniques. Electron transfer dissociation (ETD) yields less complicated spectra, with minimum fragmentation of the AMP itself. In contrast, collision-induced dissociation (CID) and high-energy collision (HCD) fragmentation caused AMP to fragment, generating characteristic ions suitable for identification of adenylylated peptides. These ions and mass losses from the AMP group were highly dependent on the identity of the adenylylated amino acid (e.g., different reporter ions for Tyr-AMP vs Thr-AMP). Upon CID and HCD fragmentation, the entire AMP group is likely to dissociate if attached to a threonine, resulting in a loss of 347 Da. In the case of Tyr-AMP, the predominant losses are either adenine (−135 Da) or adenosine (−249 Da). Upon ETD fragmentation, AMP is fairly stable, and thereby the peptide fragmentation spectra are easier to interpret manually as well as by search engines. It was shown to be possible to identify the adenylylated peptides with search engines, exemplified for Mascot if the correct neutral losses are applied. Upon ETD

■

RELATED MODIFICATIONS Nucleotides are generally versatile high energy donors of activated functionalities and thus can modify proteins differently depending on the potential substituent at the phosphates. Despite the apparently conserved core structures and sequence motifs of Fic enzymes, it has been surprising to observe that some of these enzymes bind other nucleotides differently, resulting in unexpected PTMs. The bacterial Fic family protein Doc also binds to ATP but forces the nucleotide into an inverted orientation relative to the classical Fic domains. As a consequence, the (AG)N(GK)-anion pocket that coordinates the α-phosphate of ATP in other Fic proteins now binds to the γ-phosphate, thus leading to protein phosphorylation.12,20,21 The phosphorylated target protein is the bacterial ribosomal elongation factor EF-Tu at T382EF‑Tu, resulting in the inhibition of bacterial translation via interfering with binding to aminoacylated tRNAs. Another example for binding to a nucleotide in a noncanonical manner is the Legionella Fic protein AnkX. This Fic enzyme instead of ATP binds to cytidine diphosphate choline (CDP-choline) and causes the phosphocholination of Rab1 at S76 (Figure 4A).18,67 Structural analysis of AnkX revealed that CDP-choline (similar to ATP in Doc enzymes) is positioned in the nucleotide binding pocket in an inverted orientation, too.19 Therefore, not the α-phosphate but the βphosphate is bound in the (AG)N(GK)-anion pocket, thus causing the transfer of phosphocholine to the target protein (Figure 4B). To analyze protein phosphocholination, methods for the generation of phosphocholinated peptides at threonine, serine, and tyrosine have been derived.68 It appears as if the family of Fic enzymes can utilize different nucleotides as donors for the modification of proteins. This hypothesis makes the biology of Fic proteins and possibly related enzymes extremely exciting. However, it also complicates the search for the physiological roles and targets of a given uncharacterized Fic: Since the nucleotide substrate cannot be deduced form amino acid sequence analyses (because the Fic motif is essentially a phosphate binding G



Reviews

recognition motif. Third, adenylylation is brought about mainly by enzymes from potential or actual bacterial pathogens that interfere with biological processes of host cells; therefore, pathogenic adenylylations may also be interesting for understanding the emergence and mechanisms of infections and thus also for fighting them. Sequence homologies of the Fic family of enzymes suggest that there are thousands of potential AMP (or nucleotidyl) transferases for which the protein (and/or nucleotide) substrates are still unknown. There are, however, still many open questions remaining. The molecular consequences of protein adenylylation generally cannot be predicted, and its effect on the activities, structures, protein interaction profiles, or stabilities of the modified proteins need to be investigated individually. Even though adenylylation has been mainly recognized in the process of infection, the question remains whether this modification occurs almost exclusively via bacterial agents or whether it is a more general and currently uncharacterized PTM in eukaryotic cells. To elucidate these questions, reliable and generic methods for detecting protein adenylylation need to be developed. Recent studies on the use of ATP derivatives bearing chemical handles (e.g., alkynyl of fluorescence tags at the N6 position of the adenine of ATP) have provided the first proof of concept work in which these functionalities could be installed on protein substrates. Detailed kinetic analyses on selected AMP transferases, however, revealed that such ATP derivatives may not be generally accepted for every transferase and therefore preclude their universal application as adenylylation detection probes. Also, these ATP derivatives can be used for in vitro studies, only, since their cell delivery is difficult. Consequently, they cannot be applied directly in vivo, therefore potentially limiting the significance of the results obtained. The inherent limitations of using ATP derivatives may be overcome if protein adenylylation could be detected using high resolution MS techniques such that AMP transfer can be directly detected. Monitoring specific AMP-amino-acid fragmentation patterns as a readout for protein adenylylation has been shown to be applicable. Nevertheless, the development of standard MS protocols will depend on enrichment strategies of AMP-modified proteins. Here, Tyr-AMP, Thr-AMP, and SerAMP specific antibodies may prove valuable in the future. Even if standard protocols were available, identification of the adenylylated substrate may be difficult since AMP transferring enzymes could potentially require (unknown) stimulating partners (e.g., some Fic enzymes obviously depend on the release of inhibition by inhibitory sequences). Very interestingly, Fic-family enzymes and related proteins appear to be able to also utilize nucleotides different from ATP for protein modification. The spectrum of PTMs caused by these enzymes may therefore be broader than currently anticipated. This presumption, however, complicates the search for protein substrates since the correct nucleotide (GTP, CDPcholine, UDP-glucose, etc.) needs to be identified first before the protein substrates can be searched for (e.g., by using MS or PTM-specific antibodies). Therefore, the analysis of protein adenylylation and related modifications will require combined efforts of chemistry, chemical biology, structural biology, proteomics, and microbiology.

Figure 4. Modifications related to adenylylation. (A) Reaction scheme of protein phosphocholination. The Legionella enzyme AnkX utilizes CDP-choline to transfer a phosphocholine moiety to S76 of Rab1b with CMP as a byproduct. (B) Hypothesized reaction mechanism of protein phosphocholination by AnkX. AnkX contains a Fic-related sequence motif that binds the nucleotide in a reversed orientation when compared to canonical Fic proteins. The (AG)N(GK) anion pocket coordinates the β-phosphate of CDP-choline, whereas canonical Fics would bind the α-phosphate of ATP. Whether Mg2+ is also coordinated by the Fic motif is currently unknown (model based on refs 16 and 19).

element, only), the identification of modified proteins in cell lysates is dramatically hampered. MS based methods require the a priory knowledge of the transferred functionality, but since also rare or unconventional nucleotides may be utilized by the Fic protein (e.g., UDP-glucose, NAD, etc.), the prediction of the target mass is difficult.

■

CONCLUSIONS AND FUTURE DIRECTIONS Protein adenylylation is emerging as a very interesting posttranslational modification. First, the formed phosphodiester is stable under physiological conditions and therefore in principle allows enzymatic control over the protein modification state. Second, a relatively bulky compound with different chemical substituents (i.e., phosphate, ribose, and adenine) is introduced that significantly influences the accessible binding site(s) of the modified protein but also hypothetically could act as a specific

■

AUTHOR INFORMATION

Corresponding Authors

*Phone: +46 (0)70 4442772. E-mail: christian.hedberg@umu. se. H



Reviews

*Tel.: +49 89 289 13343. E-mail: [email protected].

(16) Engel, P., Goepfert, A., Stanger, F. V., Harms, A., Schmidt, A., Schirmer, T., and Dehio, C. (2012) Adenylylation control by intra- or intermolecular active-site obstruction in Fic proteins. Nature 482, 107−110. (17) Mattoo, S., Durrant, E., Chen, M. J., Xiao, J., Lazar, C. S., Manning, G., Dixon, J. E., and Worby, C. A. (2011) Comparative analysis of Histophilus somni immunoglobulin-binding protein A (IbpA) with other fic domain-containing enzymes reveals differences in substrate and nucleotide specificities. J. Biol. Chem. 286, 32834− 32842. (18) Mukherjee, S., Liu, X. Y., Arasaki, K., McDonough, J., Galan, J. E., and Roy, C. R. (2011) Modulation of Rab GTPase function by a protein phosphocholine transferase. Nature 477, 103−106. (19) Campanacci, V., Mukherjee, S., Roy, C. R., and Cherfils, J. (2013) Structure of the Legionella effector AnkX reveals the mechanism of phosphocholine transfer by the FIC domain. EMBO J. 32, 1469−1477. (20) Cruz, J. W., Rothenbacher, F. P., Maehigashi, T., Lane, W. S., Dunham, C. M., and Woychik, N. A. (2014) Doc Toxin Is a Kinase That Inactivates Elongation Factor Tu. J. Biol. Chem. 289, 7788−7798. (21) Castro-Roa, D., Garcia-Pino, A., De Gieter, S., van Nuland, N. A., Loris, R., and Zenkin, N. (2013) The Fic protein Doc uses an inverted substrate to phosphorylate and inactivate EF-Tu. Nat. Chem. Biol. 9, 811−817. (22) Aktories, K. (2011) Bacterial protein toxins that modify host regulatory GTPases. Nat. Rev. Microbiol. 9, 487−498. (23) Cherfils, J., and Zeghouf, M. (2013) Regulation of Small GTPases by GEFs, GAPs, and GDIs. Physiol. Rev. 93, 269−309. (24) Schoebel, S., Cichy, A. L., Goody, R. S., and Itzen, A. (2011) Protein LidA from Legionella is a Rab GTPase supereffector. Proc. Natl. Acad. Sci. U. S. A. 108, 17945−17950. (25) Gazdag, E. M., Streller, A., Haneburger, I., Hilbi, H., Vetter, I. R., Goody, R. S., and Itzen, A. (2013) Mechanism of Rab1b deactivation by the Legionella pneumophila GAP LepB. EMBO Rep. 14, 199−205. (26) Oesterlin, L. K., Goody, R. S., and Itzen, A. (2012) Posttranslational modifications of Rab proteins cause effective displacement of GDP dissociation inhibitor. Proc. Natl. Acad. Sci. U. S. A. 109, 5621−5621. (27) Hardiman, C. A., and Roy, C. R. (2014) AMPylation Is Critical for Rab1 Localization to Vacuoles Containing Legionella pneumophila. mBio 5, e01035−01013. (28) Woolery, A. R., Yu, X., LaBaer, J., and Orth, K. (2014) AMPylation of Rho GTPases subverts multiple host signaling processes. J. Biol. Chem. 289, 32977−32988. (29) Ham, H., Woolery, A. R., Tracy, C., Stenesen, D., Kramer, H., and Orth, K. (2014) Unfolded protein response-regulated dFic reversibly AMPylates BiP during endoplasmic reticulum homeostasis. J. Biol. Chem., DOI: 10.1074/jbc.M1114.612515. (30) Rahman, M., Ham, H. L., Liu, X. R., Sugiura, Y., Orth, K., and Kramer, H. (2012) Visual neurotransmission in Drosophila requires expression of Fic in glial capitate projections. Nat. Neurosci. 15, 871− 875. (31) Bukau, B., Weissman, J., and Horwich, A. (2006) Molecular chaperones and protein quality control. Cell 125, 443−451. (32) Stadtman, E. R., Shapiro, B. M., Kingdon, H. S., Woolfolk, C. A., and Hubbard, J. S. (1968) Cellular regulation of glutamine synthetase activity in Escherichia coli. Adv. Enzyme Regul. 6, 257−289. (33) Kingdon, H. S., Shapiro, B. M., and Stadtman, E. R. (1967) Regulation of glutamine synthetase, VIII. ATP: glutamine synthetase adenylyltransferase, an enzyme that catalyzes alterations in the regulatory properties of glutamine synthetase. Proc. Natl. Acad. Sci. U. S. A. 58, 1703−1710. (34) Hart, G. W., Slawson, C., Ramirez-Correa, G., and Lagerlof, O. (2011) Cross talk between O-GlcNAcylation and phosphorylation: roles in signaling, transcription, and chronic disease. Annu. Rev. Biochem. 80, 825−858. (35) Yu, X., Woolery, A. R., Luong, P., Hao, Y. H., Grammel, M., Westcott, N., Park, J., Wang, J., Bian, X., Demirkan, G., Hang, H. C., Orth, K., and LaBaer, J. (2014) Copper-catalyzed azide-alkyne

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors would like to acknowledge funding by the priority program 1623 (SPP1623) of the German research foundations (DFG). A.I. is grateful to support by a DFG-funded collaborative research center (SFB1035, project B05). C.H. thanks the Knut and Alice Wallenberg foundation for generous support.

■

REFERENCES

(1) Stadtman, E. R. (2001) The story of glutamine synthetase regulation. J. Biol. Chem. 276, 44357−44364. (2) Müller, M. P., Peters, H., Blümer, J., Blankenfeldt, W., Goody, R. S., and Itzen, A. (2010) The Legionella Effector Protein DrrA AMPylates the Membrane Traffic Regulator Rab1b. Science 329, 946− 949. (3) Yarbrough, M. L., Li, Y., Kinch, L. N., Grishin, N. V., Ball, H. L., and Orth, K. (2009) AMPylation of Rho GTPases by Vibrio VopS disrupts effector binding and downstream signaling. Science 323, 269− 272. (4) Worby, C. A., Mattoo, S., Kruger, R. P., Corbeil, L. B., Koller, A., Mendez, J. C., Zekarias, B., Lazar, C., and Dixon, J. E. (2009) The fic domain: regulation of cell signaling by adenylylation. Mol. Cell 34, 93− 103. (5) Müller, M. P., Albers, M. F., Itzen, A., and Hedberg, C. (2014) Exploring Adenylylation and Phosphocholination as Post-Translational Modifications. Chembiochem 15, 19−26. (6) Xu, Y., Carr, P. D., Vasudevan, S. G., and Ollis, D. L. (2010) Structure of the adenylylation domain of E. coli glutamine synthetase adenylyl transferase: evidence for gene duplication and evolution of a new active site. J. Mol. Biol. 396, 773−784. (7) Xu, Y., Zhang, R., Joachimiak, A., Carr, P. D., Huber, T., Vasudevan, S. G., and Ollis, D. L. (2004) Structure of the N-terminal domain of Escherichia coli glutamine synthetase adenylyltransferase. Structure 12, 861−869. (8) Holm, L., and Sander, C. (1995) DNA polymerase β belongs to an ancient nucleotidyltransferase superfamily. Trends Biochem. Sci. 20, 345−347. (9) Gavriljuk, K., Schartner, J., Itzen, A., Goody, R. S., Gerwert, K., and Kotting, C. (2014) Reaction Mechanism of Adenylyltransferase DrrA from Legionella pneumophila Elucidated by Time-Resolved Fourier Transform Infrared Spectroscopy. J. Am. Chem. Soc. 136, 9338−9345. (10) Müller, M. P., Shkumatov, A. V., Oesterlin, L. K., Schoebel, S., Goody, P. R., Goody, R. S., and Itzen, A. (2012) Characterization of Enzymes from Legionella pneumophila Involved in Reversible Adenylylation of Rab1 Protein. J. Biol. Chem. 287, 35036−35046. (11) Kinch, L. N., Yarbrough, M. L., Orth, K., and Grishin, N. V. (2009) Fido, a novel AMPylation domain common to fic, doc, and AvrB. PLoS One 4, e5818. (12) Garcia-Pino, A., Zenkin, N., and Loris, R. (2014) The many faces of Fic: structural and functional aspects of Fic enzymes. Trends Biochem. Sci. 39, 121−129. (13) Luong, P., Kinch, L. N., Brautigam, C. A., Grishin, N. V., Tomchick, D. R., and Orth, K. (2010) Kinetic and Structural Insights into the Mechanism of AMPylation by VopS Fic Domain. J. Biol. Chem. 285, 20155−20163. (14) Xiao, J. Y., Worby, C. A., Mattoo, S., Sankaran, B., and Dixon, J. E. (2010) Structural basis of Fic-mediated adenylylation. Nat. Struct. Mol. Biol. 17, 1004−1010. (15) Bunney, T. D., Cole, A. R., Broncel, M., Esposito, D., Tate, E. W., Katan, M. (2014) Crystal Structure of the Human, FIC-Domain Containing Protien HYPE and Implications for Its Functions, Structure 22, DOI: 10.1016/j.str.2014.1010.1007. I



Reviews

cycloaddition (click chemistry)-based Detection of Global Pathogenhost AMPylation on Self-assembled Human Protein Microarrays. Mol. Cell. Proteomics 13, 3164−3176. (36) Feng, F., Yang, F., Rong, W., Wu, X., Zhang, J., Chen, S., He, C., and Zhou, J. M. (2012) A Xanthomonas uridine 5′-monophosphate transferase inhibits plant immune kinases. Nature 485, 114−118. (37) Goepfert, A., Stanger, F. V., Dehio, C., and Schirmer, T. (2013) Conserved Inhibitory Mechanism and Competent ATP Binding Mode for Adenylyltransferases with Fic Fold. PLoS One 8, e64901. (38) Isberg, R. R., O’Connor, T. J., and Heidtman, M. (2009) The Legionella pneumophila replication vacuole: making a cosy niche inside host cells. Nat. Rev. Microbiol. 7, 13−24. (39) Neunuebel, M. R., Chen, Y., Gaspar, A. H., Backlund, P. S., Yergey, A., and Machner, M. P. (2011) De-AMPylation of the Small GTPase Rab1 by the Pathogen Legionella pneumophila. Science 333, 453−456. (40) Tan, Y. H., and Luo, Z. Q. (2011) Legionella pneumophila SidD is a deAMPylase that modifies Rab1. Nature 475, 506−509. (41) Chen, Y., Tascon, I., Neunuebel, M. R., Pallara, C., Brady, J., Kinch, L. N., Fernandez-Recio, J., Rojas, A. L., Machner, M. P., and Hierro, A. (2013) Structural Basis for Rab1 De-AMPylation by the Legionella pneumophila Effector SidD. PLoS Pathogens 9, e1003382. (42) Ingmundson, A., Delprato, A., Lambright, D. G., and Roy, C. R. (2007) Legionella pneumophila proteins that regulate Rab1 membrane cycling. Nature 450, 365−369. (43) Jaggi, R., vanHeeswijk, W. C., Westerhoff, H. V., Ollis, D. L., and Vasudevan, S. G. (1997) The two opposing activities of adenylyl transferase reside in distinct homologous domains, with intramolecular signal transduction. EMBO J. 16, 5562−5571. (44) van Heeswijk, W. C., Rabenberg, M., Westerhoff, H. V., and Kahn, D. (1993) The genes of the glutamine synthetase adenylylation cascade are not regulated by nitrogen in Escherichia coli. Mol. Microbiol. 9, 443−457. (45) Jiang, P., and Ninfa, A. J. (2009) Reconstitution of Escherichia coli glutamine synthetase adenylyltransferase from N-terminal and Cterminal fragments of the enzyme. Biochemistry 48, 415−423. (46) Jiang, P., Pioszak, A. A., and Ninfa, A. J. (2007) Structurefunction analysis of glutamine synthetase adenylyltransferase (ATase, EC 2.7.7.49) of Escherichia coli. Biochemistry 46, 4117−4132. (47) Brown, M. S., Segal, A., and Stadtman, E. R. (1971) Modulation of glutamine synthetase adenylylation and deadenylylation is mediated by metabolic transformation of the P II -regulatory protein. Proc. Natl. Acad. Sci. U. S. A. 68, 2949−2953. (48) Mangum, J. H., Magni, G., and Stadtman, E. R. (1973) Regulation of glutamine synthetase adenylylation and deadenylylation by the enzymatic uridylylation and deuridylylation of the PII regulatory protein. Arch. Biochem. Biophys. 158, 514−525. (49) Anderson, W. B., and Stadtman, E. R. (1970) Glutamine synthetase deadenylylation: A phosphorolytic reaction yiedling ADP as nucleotide product. Biochem. Biophys. Res. Commun. 41, 704−709. (50) Smit, C., Blümer, J., Eerland, M. F., Albers, M. F., Muller, M. P., Goody, R. S., Itzen, A., and Hedberg, C. (2011) Efficient Synthesis and Applications of Peptides containing Adenylylated Tyrosine Residues. Angew. Chem., Int. Ed. Engl. 50, 9200−9204. (51) Albers, M. F., van Vliet, B., and Hedberg, C. (2011) Amino Acid Building Blocks for Efficient Fmoc Solid-Phase Synthesis of Peptides Adenylylated at Serine or Threonine. Org. Lett. 13, 6014−6017. (52) Ogura, K., Shigenaga, A., Ebisuno, K., Hirakawa, H., and Otaka, A. (2012) Fmoc-based solid-phase synthesis of adenylylated peptides using diester-type adenylylated amino acid derivatives. Tetrahedron Lett. 53, 3429−3432. (53) Grammel, M., and Hang, H. C. (2013) Chemical reporters for biological discovery. Nat. Chem. Biol. 9, 475−484. (54) Heal, W. P., and Tate, E. W. (2010) Getting a chemical handle on protein post-translational modification. Org. Biomol. Chem. 8, 731− 738. (55) Grammel, M., Hang, H., and Conrad, N. K. (2012) Chemical Reporters for Monitoring RNA Synthesis and Poly(A) Tail Dynamics. Chembiochem 13, 1112−1115.

(56) Grammel, M., Luong, P., Orth, K., and Hang, H. C. (2011) A Chemical Reporter for Protein AMPylation. J. Am. Chem. Soc. 133, 17103−17105. (57) Lewallen, D. M., Steckler, C. J., Knuckley, B., Chalmers, M. J., and Thompson, P. R. (2012) Probing adenylation: using a fluorescently labelled ATP probe to directly label and immunoprecipitate VopS substrates. Mol. Biosyst 8, 1701−1706. (58) Chung, H. K., and Rhee, S. G. (1984) Separation of GlutamineSynthetase Species with Different States of Adenylylation by Chromatography on Monoclonal Anti-Amp Antibody-Affinity Columns. Proc. Natl. Acad. Sci. Biol. 81, 4677−4681. (59) Meyer, T., and Hilz, H. (1986) Production of Anti-(AdpRibose) Antibodies with the Aid of a Dinucleotide-PyrophosphataseResistant Hapten and Their Application for the Detection of Mono(Adp-Ribosyl)Ated Polypeptides. Eur. J. Biochem. 155, 157−165. (60) Hao, Y. H., Chuang, T., Ball, H. L., Luong, P., Li, Y., FloresSaaib, R. D., and Orth, K. (2010) Characterization of a rabbit polyclonal antibody against threonine-AMPylation. J. Biotechnol. 151, 251−254. (61) Zhao, Y. M., and Jensen, O. N. (2009) Modification-specific proteomics: Strategies for characterization of post-translational modifications using enrichment techniques. Proteomics 9, 4632−4641. (62) Petinaki, E., Guerin-Faublee, W., Pichereau, V., Villers, C., Achard, A., Malbruny, B., and Leclercq, R. (2008) Lincomycin resistance gene lnu(D) in Streptococcus uberis. Antimicrob. Agents Chemother. 52, 626−630. (63) Achard, A., Villers, C., Pichereau, V., and Leclercq, R. (2005) New lnu(C) gene conferring resistance to lincomycin by nucleotidylation in Streptococcus agalactiae UCN36. Antimicrob. Agents Chemother. 49, 2716−2719. (64) Li, Y., Al-Eryani, R., Yarbrough, M. L., Orth, K., and Ball, H. L. (2011) Characterization of AMPylation on Threonine, Serine, and Tyrosine Using Mass Spectrometry. J. Am. Soc. Mass Spectrom. 22, 752−761. (65) Hansen, T., Albers, M., Hedberg, C., and Sickmann, A. (2013) Adenylylation, MS, and proteomics-Introducing a “new” modification to bottom-up proteomics. Proteomics 13, 955−963. (66) Lewallen, D. M., Sreelatha, A., Dharmarajan, V., Madoux, F., Chase, P., Griffin, P. R., Orth, K., Hodder, P., and Thompson, P. R. (2014) Inhibiting AMPylation: A Novel Screen To Identify the First Small Molecule Inhibitors of Protein AMPylation. ACS Chem. Biol. 9, 433−442. (67) Itzen, A., and Goody, R. S. (2011) Covalent Coercion by Legionella pneumophila. Cell Host Microbe 10, 89−91. (68) Albers, M. F., and Hedberg, C. (2013) Amino Acid Building Blocks for Fmoc Solid-Phase Synthesis of Peptides Phosphocholinated at Serine, Threonine, and Tyrosine. J. Org. Chem. 78, 2715−2719.

J


Molecular Perspectives on Protein Adenylylation - ACS Chemical

Recommend Documents