Approaches to Study Phosphatases - ACS Publications - American

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Approaches to Study Phosphatases Sara Fahs,† Pablo Lujan,† and Maja Köhn*,† †

European Molecular Biology Laboratory, Genome Biology Unit, Meyerhofstrasse 1, 69117 Heidelberg, Germany S Supporting Information *

ABSTRACT: Phosphatases play key roles in normal physiology and diseases. Studying phosphatases has been both essential and challenging, and the application of conventional genetic and biochemical methods has led to crucial but still limited understanding of their mechanisms, substrates, and exclusive functions within highly intricate networks. With the advances in technologies such as cellular imaging and molecular and chemical biology in terms of sensitive tools and methods, the phosphatase field has thrived in the past years and has set new insights for cell signaling studies and for therapeutic development. In this review, we give an overview of the existing interdisciplinary tools for phosphatases, give examples on how they have been applied to increase our understanding of these enzymes, and suggest how theyand other tools yet barely used in the phosphatase fieldmight be adapted to address future questions and challenges.

O

(ii) As a result of the evolutionary diversity, the lower numbers of phosphatase genes compared to that of kinase genes6−9 (iii) The different and complex regulation of the diverse phosphatase families, adding a layer of complexity which results in an equal number of phosphatases compared to kinases at the protein level (see phosphoprotein phosphatases in section 1.1)6−9 (iv) Their intricate substrate profiles, as some phosphatases can recognize proteins, lipids, or carbohydrates and some phosphatases require regulatory proteins for substrate recognition, which makes substrate prediction in many cases an impossible task7−9 (v) The detection of a negative signal (removal of a phosphate group) that needs to be placed there first (phosphorylation) before it can be recognized by a phosphatase for dephosphorylation3,7 (vi) The highly transient interaction between most phosphatases and their substrates3 (vii) The often critical need for a specific context when studying a phosphatase of interest due to point v)7,9,10 (viii) The difficulties in designing specific modulators of phosphatase activity; for instance, the overall conserved catalytic site of phosphatases lacks a defined pocket for a small molecule

ne of the most common mechanisms by which gene product function is tightly regulated in cellular systems is the reversible phosphorylation of proteins and other cellular molecules, which enables a highly dynamic and coordinated regulation of cellular processes.1 The phosphorylation balance is crucial, as its aberrant regulation is known to lead to diseases.2,3 Since kinases and phosphatases, which respectively add and remove phosphate groups on protein or nonprotein substrates, are often simultaneously active, and no new proteins need to be synthesized, reversible phosphorylation is rapid, and it has usually a high turnover rate.3 The fact that protein kinases and phosphatases (ptases) constitute 2−4% of the genes in a typical eukaryotic genome underlines that protein phosphorylation is an ancient, universal, and crucial means for the regulation of cell physiology.1,4 Since both kinases and phosphatases equally contribute to balancing phosphorylation levels, they pose as equally attractive targets to study. Nevertheless, while strong progress has been achieved, our understanding of phosphatases still lags behind that of kinases. The general notion is that studying phosphatases is highly challenging, and a persistent misconception is that phosphatases are less specific and less tightly regulated than kinases.3 There are several underlying reasons for this, such as the following: (i) Their high evolutionary diversity and complexity based on different ancestors (i.e., evolutionary unrelated) compared to the evolution of kinases based on a common ancestor5−7 © XXXX American Chemical Society

Received: July 1, 2016 Accepted: September 12, 2016

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DOI: 10.1021/acschembio.6b00570 ACS Chem. Biol. XXXX, XXX, XXX−XXX

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sequence DxDx(V/T) (x = any amino acid), and it contains enzymes with diverse substrate specificity such as toward pSer/ Thr/Tyr, nucleotides, and pyridoxal-5′phosphate.4,6,15 The protein tyrosine phosphatase (PTP) family, defined by its catalytic signature Cx5R, is known for its diversity in domain structure and substrate preference. Contrary to the traditional PSTPs, the class I PTPs are evolutionarily related.6,17 This class consists of nontransmembrane PTPs and receptor-like PTPs, which are both largely pTyr-specific, as well as the dualspecificity phosphatases (DSPs or DUSPs), which show diverse substrate specificity dephosphorylating not only pTyr containing proteins but also pSer/Thr, carbohydrates, mRNA, and phosphoinositides.8,17,18 Further nonevolutionary related phosphatase families include lipid and nucleotide hydrolases.16 However, the definition of these families has been less clear, and different classifications have been used in the past.6,16 For example, some SAC domain family members have been grouped with the PTPs7 or within a lipid phosphatase superfamily into the inositol phosphate 5-phosphatase (INPP5) subfamily16 or grouped as a separate family due to missing protein structure classification data in a work which used a new structure-sequence-based approach to reclassify human phosphatases,6 which is the most recent to have been updated.19 Such classification discrepancies are also found, albeit to a lesser extent, within the HAD, PPM, and PTP superfamilies and are due to different criteria and bioinformatics approaches used.6,7,16,19 Furthermore, new perspectives on how to view and classify phosphatases, for example the PTPs, are currently being suggested in order to help understand their biology.20 Together, this highlights not only the complexity of phosphatases but also the lack of understanding of a large number of the phosphatases, which extrapolates to the lack of knowledge of their substrate specificities and roles.6 For an in depth description of the different phosphatase families, the reader is referred to previous reviews and references therein.6−9,13,15,16,21 To conclude, the classification of phosphatases has been and is still being refined as new information on phosphatases becomes available and new bioinformatics methods are developed, and this has been an active area in phosphatase research in recent years. 1.2. Resources for Phosphatases. Whereas for many years there have been computational resources for kinases such as databases and substrate prediction tools,5,22,23 comprehensive resources dedicated to all phosphatases were more recently introduced.6,16 In fact, even the Universal Protein Database (UniProt)24 has in the past reported on the kinases that phosphorylate certain sites on proteins but has largely neglected naming the phosphatases that dephosphorylate specific sites. By now, there are several resources available that we introduce in this section. Table 1 lists the online resources discussed here. Phosphatases, just like other proteins, can be found in “general” databases. The most commonly used include the above-mentioned UniProt24 and the Human Protein Reference Database (HPRD).25 Another helpful protein database for phosphatase searching is the Research Collaboratory for Structural Bioinformatics Protein Data Bank (RSCB PDB), which is a protein structure database; it contains 3D biological macromolecular structures, which were obtained either by X-ray crystallography or by nuclear magnetic resonance (NMR) spectroscopy.26 However, the identification of specific substrates and (de)phosphorylation sites is essential for understanding the molecular signaling mechanisms. Computational tools such as

(such as ATP for kinases), which can be used as a starting point for the screening or design of lead compounds, and the preference for negatively charged compounds in the active site of phosphatases imposes problems in bioavailability2,3. Still, the importance of phosphatases in both physiological regulation and serious pathological conditions encompassing malignancies, diabetes, asthma, cystic fibrosis, immunosuppression, or cardiovascular diseases3,11 grants them an importance that they seem to have once lost in the shadows of the spotlight of the kinases. While general genetic and biochemical strategies like knockdowns have widely contributed to the understanding of phosphatases, such methods also have limitations in informing on rapid kinetic changes, elucidating functional effects of specific phosphorylation events, and linking phosphatase activity to cellular localization.12 To overcome such restraints, a range of technologies have made their way into phosphatase research during the past decade and have been applied to enlighten missing details in the mechanisms of phosphatase networks. It is now clear that the actual differences between phosphatases in terms of structure and mechanism breed differences in the methods used to study them. In this review, we address biological questions concerning phosphatases and how to go about them, giving an overview of the available and most recent tools to study them, including resources and experimental methods at the heart of chemistry and biology. We discuss how these have been used to increase our understanding of phosphatases and how they and other new useful methods that have barely been used in phosphatase research might be applied to address future questions and challenges. A comprehensive list of abbreviations and a glossary for a more detailed description of the methods is given in the Supporting Information to enable the inclusion of multiple methods in a concise manner. 1. A Brief Guide to the Classification and Resources of Phosphatases and Dephosphorylation Sites. 1.1. Classification of Phosphatases. Historically, protein phosphatases have been grouped according to their substrate specificity and distinct catalytic mechanisms into the protein serine/ threonine phosphatases (PSTPs or PSPs) and the protein tyrosine phosphatases (PTPs), as detailed in two excellent historical perspectives on these proteins.7,9 These broad and common classes were further classified into “superfamilies”.8,13 For the PSTPs, these were phosphoprotein phosphatases (PPPs, such as PP1) and the metal-dependent protein phosphatases (PPMs, also called PP2Cs), which account for the majority of phosphoserine (pSer) and phospho-threonine (pThr) dephosphorylation.4,13 Both have a metal-dependent catalytic mechanism but are unrelated regarding their sequence.9,13,14 Whereas the PPMs include multidomain proteins, the PPPs are characterized by the formation of holoenzymes through binding of their catalytic subunit to regulatory proteins.9 These regulatory proteins define substrate specificity, modulate the activity, and regulate the localization of the PPPs9 and thus increase the number of the 13 phosphatase genes to hundreds of highly regulated and specific holoenzymes at the protein level. The aspartate-based FCP (TFIIF-associating component of RNA polymerase II CTD) and SCP (small CTD) phosphatases were originally grouped with the PPMs and PPPs;13 however, newer classifications based on sequence and protein fold placed this group into the more recently defined superfamily of haloacid dehalogenase (HAD) phosphatases.6,7,15,16 This superfamily is characterized by a Rossmann-like fold and the active site B


C

a

modification site properties and their biological function

Phosphosites Resources

Human Dephosphorylation Database (DEPOD) PTP database PTP central

human phosphatases; fully searchable (protein and nonprotein substrates, phosphatases, pathways, dephosphorylation sites), visualization of phosphatase-substrate relationships, links to kinase and ChEMBL DBs specific for PTPs, tyrosine-specific sequences; structures, disease association relies on genomic scale PTP prediction tool; sequence and phylogenetic analyses; relationships with other PTPs across genomes; structures; genetic studies, disease association

human phosphatases and some orthologs, information on expression profiles, substrates, interactions, and, structures

ptp.cshl.edu www.ptp-central. org

nonredundant modification sites (phosphorylation sites)

Phosphosite Plus

comprehensive, curated from scientific literature and public databases

www.rcsb.org gps.biocuckoo. org www. phosphosite. org phosphat.unihohenheim.de phosida.org www.p3db.org www. phosphopep. org www. phosphogrid. org ekpd.biocuckoo. org hupho. uniroma2.it www.depod.org

protein structure, X-ray and NMR data, relationships to sequence, functions, and diseases phosphorylation sites with cognate kinases, large-scale prediction

Eukaryotic Protein Kinase and Protein Phosphatase Database (EKPD) Human Phosphatase (HuPho)

www.uniprot.org www.hprd.org

protein sequence and annotation data for species, genes, and PTMs PPI, domain architecture, PTMs, interaction networks, disease association

URL

UniprotKB/Swiss-Prot Human Protein Reference Database (HPRD) Protein Data Bank (PDB) GPS 2.0

main characteristics

References a, ref 24; b, ref 25; c, ref 26; d, ref 23; e, ref 201; f, ref 31; g, ref 16; h, ref 19; i, ref 32; j, ref 33.

PTP specific protein resources

general phosphatase resources

phosphosite-specific protein resources

general protein resources

online resource

Table 1. Online Databases for Phosphatases

i j

h

g

f

e

c d

a b

referencea

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ACS Chemical Biology GPS,23 DISPHOS,27 NetPhos,28 NetPhosYeast,29 and GANNPhos30 (Table 1) serve to predict general phosphorylation sites and can be used as a starting point for phosphatase substrate research. For example, if a kinase is known for a specific phosphorylation site, this knowledge could narrow down the possibilities of specific phosphatases that could potentially dephosphorylate that residue in turn.6 A summary of useful phosphorylation Web sites can also be found on the GPS Web site (gps.biocuckoo.org). As a specific database on protein phosphatases and protein kinases, the Eukaryotic Protein Kinase and protein Phosphatase Database (EKPD) is a comprehensive resource including to date 50 433 protein kinases and 11 296 protein phosphatases (excluding nonprotein phosphatases) from eukaryotic organisms.31 Two online databases focus on human phosphatases. The Human Phosphatase (HuPho) Web portal includes Ser, Thr, and Tyr phosphatases, as well as hydrolases; is searchable for 197 phosphatases and PPP regulatory proteins; and provides information on expression profiles, substrates, interactions, structures, and orthologues.16 The human DEPhOsphorylation Database (DEPOD) is a manually curated resource on 237 human phosphatases including 11 inactive ones; searchable for phosphatases, substrates (nonprotein and protein), pathways, and dephosphorylation sites; and containing a sequence search tool for interrogating relationships with proteins from other organisms.6,19 DEPOD also contains an interactive network of human phosphatases and their substrates.6,19 As a more focused database, the PTP database offers access to sequences, multiple sequence alignments, phylogenetic trees, structures and structural alignments, and links to human diseases.32 More recently, PTP central was established as a comprehensive resource for PTPs, which relies on a genomic scale PTP prediction tool termed Y-Phosphatomer, making available diverse information on PTPs in health and disease, as well as PTP sequence predictions and multiple sequence and phylogenetic analyses online.33 Such bioinformatics tools enable broad access to extensive and curated information to experts and nonexperts of phosphatases, which strongly supports their research and, in the long run, will help deepen our understanding of phosphatases and their networks. 2. Identification of Phosphatase Substrates. A total of 226 human active phosphatases have been identified, 194 of which have at least one substrate identified amounting to a total of 305 protein and 89 nonprotein substrates, according to the DEPOD database.19 For comparison, the kinase database RegPhos lists 10 257 protein substrates,34 illustrating the knowledge gap between these enzymes families. Thus, substrate identification is still one of the main challenges in the phosphatase field, for reasons which include the intricate substrate profiles, the regulatory elements that phosphatases sometimes use for substrate recognition, the detection of a negative signal, and the highly transient interaction between most phosphatases and their substrates (see the Introduction).3,7−9,21,35 However, new approaches have greatly helped to overcome these issues: In this section, we collect different MS- and intact cell-based approaches used successfully to identify phosphatase protein and nonprotein substrates (enzymatic approaches, which are also briefly mentioned in section 3.1, and bioinformatics methods are described in detail elsewhere6,36−41). Most of the following techniques can also be used for studying protein− protein interactions (PPIs). Of note, for a more detailed description of the methods and the abbreviations, the reader

is referred to the glossary in the Supporting Information. Supporting Table S1 lists all methods in detail, including advantages and limitations. 2.1. MS-based Methods for Protein Substrate Identification. Generally, phosphatase substrates that were isolated from the bulk of proteins by coprecipitation using the phosphatase of interest as bait are characterized by MS. Over the past few years, different methods have been developed to improve the selectivity and sensitivity of this strategy. 2.1.1. Protein−Substrate Interaction Stabilization. Compared to other PPIs, phosphatase−substrate interactions are transient and weak, and their simple isolation is rather unlikely.40 Thus, stabilization of this interaction should be realized previously to its isolation. In the context of PTPs, substrate-trapping mutants can be used for this purpose. This principle is based on the ability of site-directed mutated PTPs to retain their substrate specificity but abolish their phosphatase activity.42−44 Mutation of the catalytic cysteine to serine (C−S) or the catalytically important aspartate from the WPD loop to alanine (D−A) were first characterized and serve as the most common substrate-trapping mutants.42−44 These two mutations can be combined to increase the substrate-trapping efficiency between them or with other mutations, like the highly conserved glutamine stabilizing the water molecule which attacks the cysteine−phosphate intermediate to alanine (Q−A)45 or the tyrosine that coordinates substrate recognition to phenylalanine (Y−F).46 Substrate-trapping efficiency among these combinations varies depending on each PTP.43 Substratetrapping mutants are extensively used in combination with other techniques for protein substrate identification due to their high selectivity. Nevertheless, for DUSPs that belong to the PTPs, this strategy can be somewhat inefficient when the substrate is not a protein.47 For PPMs or PPPs, substrate-trapping mutants have unfortunately not yet been identified. Chemical cross-linking is a good alternative for phosphatase− substrate stabilization since this approach leads to covalent binding.48,49 Unfortunately, interacting proteins might also be targeted, and protein−substrate selective interactions cannot be elucidated solely by cross-linking. However, it can be useful to corroborate already proposed substrates by showing the direct interaction. Cross-linking has also been used to elucidate phosphatase interacting proteins like the identification of the PP2C−OST1 interaction that relates the phosphatase to the plant hormone abscisic acid (ABA) signaling,50 and it has been used to determine the structure of the PP2A network.51 In order to gain selectivity, the cross-linker can be integrated in the phosphatase by site-specific unnatural amino acid incorporation by amber codon suppression.52 However, although using unnatural amino acids would be more regioselective, regulators and other interacting proteins could still be cross-linked, as was the case when this technique was used to demonstrate that VHR phosphatase activity is modulated by dimerization.53 Proximity-based assays have been mainly used in intact cells where cDNA libraries are used for screening for interactions (see section 2.2). Recently, proximity-dependent BioID (Table 2 and Figure 1) has been developed combining this principle with MS readout. A biotin ligase is fused to the bait phosphatase, leading to biotinylation of substrates and interacting and neighbor proteins, which are isolated for MS analysis.54,55 Since the biotin ligation is a fast reaction, short and weak interactions can be detected. This advantage makes BioID a potential method for phosphatase substrate identification although it is not sufficient to distinguish substrates from interacting and neighbor proteins. D


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ACS Chemical Biology Table 2. Methods for Studying Phosphatase Substratesa class MS-based methods

intact cellbased methods

lipid ptase study

subclass proteomics

phosphoproteomics Y2H-based methods FRET-based methods fluorescent colocalization FCCS-based methods

examples in ptase field

methods

coverage

1. Stabilization: substrate trapping mutants, cross-linking, unnatural amino acids, BioID, SMM 2. Isolation: GST pull-down, Co-IP, TAP 3. Identification: SILAC, ICAT, iTRAQ, SRM, MRM, SWATH IMAC, MOAC, immunoaffinity, chemical modulation Y2H, PCA, MaMTH, KISS, BiFC, PLA, M-Track FRET, UC-FRET, pcFRET, FLIM

general general general general target target

a b c d e f

immunoimmobilized baits, VIP, QD-VIP

target

none

FCCS, RICS, FLCS

target

g

labeled PIPs, ESI-MS, (MALDI-IM)-MS, (FRET) lipid binding domains, biotinylated lipids, metabolically stabilized and cross-linkable lipids, chemical-inducible dimerization

target

h

a

References: a, refs 42−46, 50, 51, 53, 57−60; b, refs 40, 82; c, refs 40, 51, 80, 81; d, refs 89−92; e, refs 94−98; f, refs 100, 101; g, refs 106, 107; h, refs 126−129, 131.

2.1.2. Substrate Isolation. After the phosphatase−substrate interaction is stabilized, it should be purified from the cell lysate before its identification. GST pull-down44,74 (Table 2) is a blind, brute, and versatile in vitro approach. However, in order to detect the substrate by GST pull-down, it must be kept phosphorylated in the cell lysate. To ensure this, there are different alternatives: deletion of endogenous phosphatase by, for example, knock-down,7 use of broad range inhibitors like pervanadate for PTPs,75 and okadaic acid76 or calyculin A9 for PPPs or overexpressing a promiscuous kinase like PTK or Src.44 In order to avoid these inconveniences, either the endogenous phosphatase can be used or the substrate-trapping or exogenous phosphatase equipped with commonly used tags (such as myc, HA, Flag, GFP) can be directly expressed in mammalian cells and purified together with its interacting proteins by coimmunoprecipitation.44 New genome editing technologies can be used to avoid overexpression artifacts; however, the realization of this approach is not always straightforward.77,78 This in-cell approach also allows locating the phosphatase in its cellular environment near its substrates. For further sample isolation improvement, tandem affinity purification (TAP) can be done79 (Table 2). To a greater or lesser extent, both techniques also lead to the isolation of other PPIs besides substrates. To differentiate between them, the use of competitive inhibitors like pervanadate for PTPs or calyculin A for PPPs is advisible.9,44 2.1.3. Substrate Identification. After substrate candidates are isolated, hits can be analyzed by SDS-PAGE and Western Blot for candidate corroboration or MS to yield the whole proteomic profile. Large-scale peptide MS quantification is generally achieved through labeled or label-free peptide LC separation coupled to ESI-MS. Isotopic labeling methods (SILAC, ICAT, iTRAQ) have a narrow dynamic range and high expense while label-free MS (SRM, MRM, SWATH) has a wide dynamic range besides being more selective, sensitive, and reproducible (Table 2).55 These approaches have been extensively applied in phosphatase substrate identification, network, and structure.40,51,80−82 Phosphoproteomic approaches using quantitative MS can be an alternative where phosphatase−substrate stabilization is not needed. It is based on the isolation and quantification of phosphopeptides from the cell lysate. The most common methods used for phosphopeptide enrichment are based on the ability of some metal oxides like TiO2 (MOAC)83 or multivalent cations like Fe(III) (IMAC)84 to form complexes

Furthermore, similarly to small molecule microarrays, phosphopeptides can be immobilized to a solid matrix or membrane and used for high-throughput protein−peptide interaction screening.56 As a readout, tagged substrate-trapping mutants57,58 can be used as wells as phosphopeptide dephosphorylation measurement59,60 allowing kinetic measurement and the study of the phosphatase regulation.59 These microarrays are useful for substrate identification and motif recognition from the phosphopeptide substrate point of view and have been developed for both PTPs57,58,60 and PSTPs59 and have been applied for example to identify protein substrates for PTP1B61 and DEP-1.62 Moreover, diverse cross-linkable or nonhydrolizable phosphotyrosine mimetics have been developed to profile phosphatase activity in complex proteomes (see section 3.1).63−66 Among them, the unnatural amino acid 2-fluoromethyl pTyr (2-FMPT) presents N- and C-termini that can be used for peptide synthesis. Different peptide sequences containing 2-FMPT have been used to generate activity-based fingerprints to study the peptide specificity for different PTPs.67 In this context, these phosphoamino acid analogs can also be incorporated in the complete substrate for phosphatase− substrate stabilization in order to further study the interaction, for example by crystallography. There are several methods to incorporate these mimetics including total chemical synthesis with a 60 amino acid length limitation, semisynthesis through the ligation between a C-terminal protein thioester and a N-Cys-containing synthetic peptide including the phosphomimetic, cysteine modification to generate a pSer mimic, Staudinger ligation to create a pTyr mimic, and site-specific unnatural amino acid incorporation.68 Compared to phosphorylation on serine, threonine, and tyrosine, phosphorylation on lysine, histidine, and cysteine is a more unstable modification and has been challenging to detect.69−71 New approaches to address the synthesis of these phosphorylated residues, for example to use them for in vitro studies on phosphatase substrate specificity or as standards in MS-based methods, include the Staudinger-phosphite reaction on lysine,70 the replacement of an activated cysteine disulfide with a phosphite reagent,71 and the chemical synthesis of phospho-histidine (pHis) analogs.72 These nonhydrolizable pHis analogs have been useful to create anti-pHis antibodies,72 which can be applied to enrich for pHis peptides for MS detection.69 By now, antibodies have been developed that can distinguish between the phosphorylation at the 1- or 3-position on pHis.73 E


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Figure 1. Methods for phosphatase substrate identification. Schematic representation of selected methods for studying phosphatase substrates: (a) BioID is a technique based on the biotin ligation by a promiscuous mutated biotin ligase BirA R118G fused to the bait phosphatase to proximal proteins within a radius of 20−30 nm. Afterward, biotinylated interaction partners are isolated by streptavidin affinity purification. (b) BiFC is a proximity-based assay where bait and prey are fused each to one-half of a fluorescent protein, leading to fluorescence upon interaction. (c) FLIMFRET takes advantage of the photophysical phenomenon where, in close proximity, a donor florophore can transfer the excited state energy to an acceptor. Bait and prey proteins are tagged with these fluorophores, respectively. This technique requires that the donor’s emission and the acceptor’s absorption spectra overlap. To overcome FRET limitations as described in the text, the fluorescence lifetime, defined as the time an electron spends in the excited state, can be measured (FLIM). (d) Fluorescently labeled baits are immuno-immobilized in the living cell to a glass surface in a patched pattern through a transmembrane domain. PPIs are recognized by colocalization of fluorescently labeled preys with the corresponding bait patch like in the example picture representation of a possible result (below) after TIRF microscopy readout. (e) Chemically induced heterodimerization where type IV 5′-ptase (5′Ptase) is recruited to the plasma membrane upon rapamycin-induced FRB-FKBP12 dimerization. PI(4,5)P2 is dephosphorylated and depleted from plasma membrane. As a consequence, a fluorescently labeled PI(4,5)P2 selective sensor (PH) cannot bind the plasma membrane and locates to the cytosol. F


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FLIM-FRET, which measures the fluorescence lifetime of the donor since that is shortened by the energy transfer to the acceptor (Figure 1).99 FLIM-FRET is by now quite commonly used in phosphatase research and has been successfully applied to corroborate interactions like PTP1B and EphA3100 or PP1 and NIPP1.101 As another FRET alternative, a protein array was developed where different fluorescently labeled baits are immunoimmobilized in the living cell to a glass surface in a patched pattern through a transmembrane domain (Figure 1). PPIs are recognized by colocalization of fluorescently labeled preys with the corresponding bait patch.102 Although it has not yet been used for phosphatase−substrate studies, it is a powerful tool since it can be used for high-throughput live cell screening as a simultaneous measure of multiple interaction partners can be assayed with only tagging the prey protein. Similarly relying on colocalization, Visual IP (VIP)103 and quantum dots-VIP (QD-VIP)104 have been developed to detect PPIs in a nonperturbing manner (Table 2). They are suitable for use in cell lysates and live cell imaging, respectively. Both have been used for kinase research and might have a similar potential for the study of phosphatases. Finally, fluorescence cross-correlation spectroscopy (FCCS)105 is a powerful method to determine that two proteins belong to the same molecular complex (Table 2). It cannot distinguish between the substrate, PPI, or if two proteins belong to the same protein complex, but FCCS has the same advantages of FRET without its limitations. Thus, it has been used in studies like the dynamics of PTPD1/EGFR-containing signaling,106 the interaction between MKP-1 and p38, and also between DUSP5 and ERK1/2.107 2.3. Methods for Nonprotein Substrate Identification. Some phosphatases have lipid or carbohydrate phosphatase activity.108−110 Laforin is the only phosphatase presenting a carbohydrate binding domain (CBD) and being able to dephosphorylate polysaccharides.36,110 However, there are more lipid phosphatases for the lipids that present hydrolyzable phosphates. In the present review, we will focus on the study of phosphatidylinositol phosphates (PIPs), as for other nonprotein substrates very few specific tools are described. PIPs represent a low percentage of the components in the cell membrane but are indispensable for many cell signaling pathways. In vitro enzymatic assays generally used for protein substrate discovery can also be used to detect lipid phosphatase activity. However, radioactive or fluorescently labeled PIPs analyzed by TLC47,111,112 or HPLC113,114 are more commonly used. Nowadays, unlabeled cell extracted lipids can also be quantified by ESI-MS115−118 or MALDI-IM-MS,115,119 but the mass overlap between different lipid species make the data analysis complex.120 For live cell imaging of the dynamics of PIP turnover, the only available technique is using fluorescently labeled protein domains that recognize specifically one of the PIP species (reviewed by Balla and Varnai121).121−123 Recently, cellpermeable smaller lipid sensors based on these domains have been developed to avoid protein overexpression.124 In any case, binding of the sensor could inhibit a lipid-mediated cellular process, and this should be taken into account when interpreting the data. Monitoring a single fluorescently labeled sensor is not per se a quantitative method. However, coexpression of the same domain labeled with CFP and YFP can be used for FRET detection on phosphoinositide patches. Dephosphorylation of

with the phosphate group. Immunoaffinity chromatography has been also used but is usually restricted to the study of PTPs due to the quality of the available antibodies.84 Different chemical modifications via β-elimination,85 cistamide linking,86 or dendrimer conjugation87 to covalently bind the phosphopeptides directly to the resin can be also used. The amount of decrease of a specific phosphopeptide under the presence of the phosphatase of interest compared to appropriate controls (inactive mutant, total protein amounts) could be an indication for a possible substrate. Unfortunately, direct substrates and downstream effects cannot be easily distinguished, and pulsechase experiments and bioinformatics analysis88 are usually necessary. Phosphoproteomics have been successfully applied for the study of the status of the phosphoproteome depending on the presence of different phosphatases like PTP1B,89 PRL3,90 PP1,91 PP2A,92 or PP6.10 2.2. Intact Cell-based Methods for Protein Substrate Identification. Defining phosphatase−substrate interactions inside living cells can provide valuable insights into signaling and is required for establishing real substrates of phosphatases. Proximity-based approaches between substrate-trapping phosphatase mutants and cDNA libraries, which is a powerful tool for high-throughput cell-based substrate screening, have been extensively used for substrate identification of PTPs. Y2H is the canonical proximity-based method for yeast cellbased PPI identification where the bait phosphatase is fused to the binding domain of a transcription factor and the cDNA library is fused to the activating domain; thus gene activation is the readout.93 Y2H has been used in the past together with substrate-trapping PTP mutants to discover PPIs and substrates like the interaction between PTP-BL with the tumor suppressor APC,94 PP1 binding to PP2A regulatory proteins,95 PTPRZ substrate candidates identification,96 and the binding of Laforin to pyruvate kinase M1/M2.97 However, despite being a robust method, Y2H has several limitations: the interaction is forced to occur in the nucleus, it is done in yeast, and transient interaction cannot be easily addressed,55 although it is commonly used in collaboration with substrate-trapping mutants. New methods based on Y2H have been developed in mammalian cells to overcome these limitations. They can be classified in three groups (Table 2): one is based on the folding of two split halves of a protein or circle-forming DNA [PCA, split TEV, BiFC (Figure 1), MaMTH, and PLA], another on an interaction-enforced enzymatic labeling reaction (M-Track, Bio-ID), and the last group on the restoration of a dysfunctional signaling pathway (KISS). Since split TEV, MaMTH, and KISS are reporter-based systems, gene activation leads to accumulation of luciferase over time and changes in the interaction cannot be measured. However, amplification of the signal makes these methods more sensitive for weak and transient interactions.55 With the exception of M-Track, which was specifically developed to detect transient phosphatase−substrate interactions and used to identify new PP2A substrates,98 most of the present techniques have been rarely used in phosphatase research. However, they are good alternatives to Y2H and have great potential for PPI and substrate identification and corroboration. FRET is a photophysical phenomenon used to determine spatiotemporal PPIs in living cells (Table 2). Although it is a technique that has been extensively used for phosphatase substrate and PPI elucidation, FRET presents some limitations like photobleaching, concentration of the fluorophore, and exposure and intensity variation. All of them can be evaded by G


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offers advantages such as handling simplicity, speed, and tunability, and particularly small molecule inhibitors can eventually be used as lead compounds for drug discovery and are required for target validation.2,134 However, only few protein phosphatase activity modulators are currently being developed for therapeutic use, and this is due to challenges such as the lack of a defined pocket in the catalytic site and the preference for negatively charged molecules in the active site (see the Introduction).3 Nevertheless, new developments in the area of chemical probes and modulators for phosphatases show that these challenges can be overcome, as we discuss in this section (for further reviews see refs 3, 11, 135, and 136). 3.1. Chemical Sensors of Phosphatase Activity. Chemical probes allow the first assessment of general phosphatase substrate activity and specificity in vitro. For example, radioactively [32P]-labeled phosphopeptides or proteins,36,37,137 31P NMR,138 chromogenic substrates like pNPP and DiFMUP (Figure 2),139 HPLC,140 and colorimetric assays36,37,141 are used to assess phosphate hydrolysis by phosphatases. Similarly to small molecule inhibitors that target the active site, phosphatasespecific probes are difficult to develop due to the conserved active site structures. Still, subtle variations such as surface potential and adjacent binding pockets enable developing specific probes.134,139 An example of such probes is an iminocoumarin derivative, which is a fluorescent probe specific for DUSPs.142 For some PTPs, selective pTyr-peptides were found.143,144 Peptide substrates incorporating phosphocoumaryl amino propionic acid (pCAP; Figure 2), a fluorogenic phosphotyrosine mimetic whose fluorescence increases upon dephosphorylation, have been used for high-throughput inhibitor screening of CD45 in T cells and macrophages.145,146 These substrates and their analogs are both specific and potent in terms of monitoring enzyme efficacy.139 Nevertheless, since these are not the natural substrate of the studied phosphatases, the kinetic parameters obtained might be different than the actual ones. To this end, phosphorylation-sensitive Sox fluorophores can be an alternative. The fluorophore is covalently appended to a cysteine residue in the substrate peptide containing the phosphotyrosine, and the dephosphorylation is monitored by a decrease of chelation-enhanced fluorescence (CHEF; Figure 2).143 The selectivity of the sensor depends on the selectivity of the peptide. Off-target activity can be decreased by target phosphatase enrichment by immunoprecipitation from the cell or tissue lysate. In addition to that, more robust fluorescence probes have been developed to avoid inaccurate results due to factors that could impact fluorescence measurements such as pH value variation and excitation light intensity: these ratiometric probes were designed for cellular imaging studies and rely on the FRET technique.147 An example includes probes with a 3′-O, 6′-O-protected fluorescein acceptor linked to a coumarin donor through a linker moiety, designed by Nagano and colleagues.147 These probes overcome the fluorescence-quenching problem that often occurs with FRET probes, using the coumarincyclohexane-fluorescein FRET cassette moiety, in which close contact of the two dyes is obstructed.139,147 The use of FRET-based reporters has enabled directly visualizing native, real-time activity of a desired phosphatase with a very high spatiotemporal resolution in living cells. An example is the calcineurin activity reporter (CaNAR) designed by Zhang and co-workers, whereby a well-studied substrate of calcineurin (NFAT) was chosen as a “dephosphorylationcompetent” molecular switch.148 The system is constructed in a

the prey lipid could be then quantified as a modulation of the FRET signal in a specific region.125 This technique has been used to monitor lipid phosphatase activity of different VSPs.126−128 For a nonradioactive in vitro kinetic study of PIP dephosphorylation, a sensor complex formed by a biotinylated lipid and its appropriate binding domain can be used. Competition between the lipid present in the sample and the biotinylated lipid for its binding domain can be detected using AlphaScreen or time-resolved FRET. This technique works in vitro but also in cell lysate, and it has been tested on PTEN’s PI(3,4,5)P3 phosphatase activity129 and to measure total PI(3,4,5)P3 cell content.130 Metabolically stabilized, photo-cross-linkable, biotinylated, or immobilized PIP derivatives have been synthesized and applied to study PIP kinases, phosphatases, and binding proteins, as reviewed previously.131 In vivo phosphatase activity modulation by protein inhibition or protein mislocalization is useful for the characterization and corroboration of phosphatase substrates. This can be accomplished by chemically induced heterodimerization, where for example the membrane localization of a lipid phosphatase like type IV 5-ptase fused to a small molecule-binding domain depends on treatment with a small molecule like rapamycin, which binds and dimerizes the phosphatase-fused domain and another domain fused to the membrane.132 Upon treatment, the type IV 5-ptase domain is localized to the plasma membrane and dephosphorylates its substrate PI(4,5)P2, which can be visualized by a loss of binding of the above-described fluorescently labeled sensor domains (Figure 1). Different systems and chemical inducers of dimerization are available for this purpose.133 Of note, given that the phosphatase activity depends on the correct orientation toward the substrate in the membrane, it may take several attempts of optimization to obtain a fusion construct that works as desired. The methods to study nonprotein phosphatases are still rather limited due to the difficult nature of the substrates and the challenging synthesis of PIP-based probes. The methods are in general rather brute and require strong phosphatase activity for detection, and currently fast changes in substrate turnover, as evident with phospholipids, are still difficult to detect in intact cells. 3. Studying the Function of Phosphatases: From Sensing to Modulation Using Tool Compounds. Given the characteristics and properties of phosphatases raised in the Introduction, studying their function is not always straightforward. Traditional genetic methods, such as knock-down approaches, have proven to be useful for the study of mainly PTP function,7 and while they have given information on some PPP (isoforms) as well, they do not provide an effective and versatile means to accurately measure protein dephosphorylation (and phosphorylation) in a temporally and spatially controlled manner.12 Knocking down a PPP affects a multitude of holoenzymes, and therefore the outcome is not generally simple to interpret.3 Potential functional redundance of related phosphatases poses another problem, and therefore traditional gene knockdowns or knockouts may not always be the best simulator of an in vivo scenario.3,134 Nevertheless, with keeping the obstacles in mind, the new genome editing technologies will also be highly valuable tools in phosphatase research, and they have been reviewed extensively elsewhere.77,78 The use of small molecules to identify cellular processes in which phosphatases are involved is a powerful complementary method to the genetic methods. The usage of tool compounds focuses mostly on modulating or sensing enzymatic activity and H


Reviews


Figure 2. Chemical tools for studying protein dephosphorylation. (a) FRET based activity sensor: a dephosphorylatable substate sandwiched between a FRET donor and an acceptor undergoes a change in fluorescence readout upon dephosphorylation by a specific phosphatase. This functions as a molecular switch that stimulates a change in the proximity and relative orientation of the dyes linked to the known substrate, leading to a FRET change. This is reversed upon rephosphorylation of the reporter by cellular kinases. (b) Endogenous phosphatase activity sensors: phosphorylation-sensitive Sox fluorophore (2D) is covalently affixed to an engineered cysteine residue in a peptide substrate. In the presence of Mg2+, the Sox fluorophore undergoes chelation-enhanced fluorescence (CHEF) when proximal to a phosphoryl-amino group. Phosphatase activity removes the phosphate group from the peptide probe, thereby reducing the affinity for Mg2+, leading to a decrease in fluorescence that can be measured. (c) Schematic representation of activity-based protein profiling: activity-based probes (ABPs) specifically label the phosphatase of interest by covalent modification of the residues essential for the catalytic mechanism. Once the phosphatase is labeled, it can be then analyzed by in-gel fluorescence scanning and/or affinity pull-down/LCMS for target identification. (d) Chemical structures of small molecule sensor (Sox, red) and substrate mimetics: phosphocoumaryl amino propionic acid (pCAP), a fluorogenic phosphotyrosine mimetic whose fluorescence increases upon dephosphorylation; 6,8- difluoro-4-methylumbelliferyl phosphate (DiFMUP); and p-nitrophenyl phosphate (pNPP), chromogenic substrates used to assess phosphate hydrolysis by phosphatases, thereby used for studying phosphatase activity.

way that conformational change resulting from dephosphorylation results in a FRET increase in response to calcineurin activity (Figure 2).148 A chemical FLIM-based sensor (see also section 2.2) was used to elucidate the mechanism underlying the spatial regulation of phosphatase activity: PTP1B was tagged with a donor chromophore protein and overexpressed in cells. To spatially sense its activity, a known pTyr-containing peptide substrate was synthesized, conjugated to an acceptor chromophore, and added to cells. Active PTP1B interacted with the substrate

resulting in a FLIM signal. Once the substrate was dephosphorylated, both the interaction and the FLIM signal were lost. This technique is useful to detect where in the cell the phosphatase is active and also to determine catalytic parameters.149 Another chemical biology technique which delivers distinctive applications to study phosphatases in complex proteomes and living cells is activity-based protein profiling (ABPP), employing probes that determine the functional states of phosphatases in complex proteomes, measure enzyme expression levels, and detect active enzymes postprocessing levels I


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ACS Chemical Biology Table 3. Examples of Modulators of Phosphatases and Their Classificationa type inhibitors of protein phosphatases

activators of protein phosphatases

1 2 3 4 5 6 7 8 9 1′ 2′ 3′ 4′ 5′ 6′ 7′

mode of action active site inhibition bidendate inhibition−binds both active site and unique adjacent peripheral site binds hydrophobic pockets adjacent to active site binds to allosteric site remote from active site−stabilizes inactive conformation disrupts subsets of holoenzymes by binding of catalytic subunit disrupts subsets of holoenzymes by binding of regulatory protein disrupts subsets of holoenzymes−targets less common docking sites inhibits substrate recruitment and/or subcellular targeting inhibition by dimerization binds catalytic domain interacts with noncatalytic, allosteric sites targets inhibitory proteins of the target phosphatase targets catalytic subunit allosterically remote from active sitedisrupts PPP holoenzyme targets catalytic subunit allosterically adjacent to active siteremoves inhibitory protein disrupts PPP holoenzymeinhibits transcription of inhibitory protein causes PTP dimerizationforms more stable dimers

phosphatase example

reference

PHLPP2 TC-PTP Scp (HAD phosphatase) PTP1B PP1/Tat PP1/GADD34 PP3 (calcineurin, PP2B) RPTPα PP2Cα PP5, SHIP1 PP1/GL complex PP1

a b c d e f g h g i j k l

PP2A

m

PP2A DEP-1

n o

a

References: a, ref 202; b, ref 203; c, ref 189; d, ref 170; e, ref 179; f, ref 180; g, ref 3; h, ref 204; i, ref 205; j, refs 196, 206; k, ref 207; l, ref 192; m, ref 193; n, ref 208; o, ref 197.

(see also section 2.1.1).150,151 These activity-based probes (ABPs) exclusively label the phosphatases of interest by covalent modification of the residue essential for the catalytic mechanism, which needs to be a nucleophilic residue like cysteine in PTPs for the principle to work. The labeled phosphatase can then be analyzed by in-gel fluorescence scanning and/or affinity pull-down/LCMS (for target identification; Figure 2).150 For PTPs, ABPs can be grouped into four different types on the basis of trapping mechanism and structure: Type I ABP is known for forming a covalent bond with a PTP (in the presence of catalytic cysteine). For instance, Zhang and colleagues have created reporter-tagged bromobenzylphosphonate probes that label many purified PTPs in an active-site-directed manner.65 Probes that belong to type II ABP contain a vinylsulfone ester of tyrosine and an azide moiety; the latter serves as a functional group for introduction of an affinity tag, resulting in a technique that overcomes low permeability issues of biotin-based probes.66 Type III and type IV ABPs share structural patterns of binding, but they differ in the key structural units: a 4-fluoromethylphenylphosphate (FMPP) group for the former152 and a 2-fluoromethylphosphotyrosine (FMPT) moiety for the latter,64 both eventually leading to specific covalent binding to PTPs. ABPs can be used for example to study the difference in phosphatase activity between healthy and diseased tissue.153 Since ABPs require a nucleophilic residue in the phosphatase for the activity-dependent trapping of the molecule, this principle cannot directly be used for other phosphatases like PPPs or PPMs, as they do not contain a nucleophile that attacks the substrate: this is done by an activated water molecule during catalysis.9,13,14 However, affinity-based probes, such as biotinylated or fluorescently labeled microcystins that bind covalently to a cysteine adjacent to the catalytic center of several PPPs,154 can be used instead for example to identify interacting proteins that do not require interactions at the catalytic site to bind to a certain PPP. 3.2. Chemical Modulators of Phosphatase Activity. This subject has been covered extensively in previous reviews, mostly for protein phosphatases.3,11,134,155−160 Here, we aim to give an overview and briefly summarize previous approaches,

mention newer developments, and add considerations for the use of phosphatase modulators. For a general classification of phosphatase modulators, the reader is referred to Table 3 (based on ref 3). Inhibitors can be considered as loss of function probes, similarly to a genetic knock-down, only more tunable.134 Finding a specific active site inhibitor for a phosphatase of interest is not devoid of difficulty (see the Introduction); nevertheless, the number of inhibitors for phosphatases is increasing steadily.134 New insights into protein structures and into crucial interactions with other proteins, the availability of large chemical libraries and various design strategies thereof, automated high throughput screens, fragment-based approaches, drug repurposing and derivation, computational docking and shape-based methods, and attempting allosteric instead of active site binding are all approaches that assist in the identification of selective inhibitors.161−166 Compound bioactivity libraries such as the ChEMBL database (www.ebi.ac.uk/chembl)167 allow scientists to quickly access information on chemical modulators of their phosphatase of interest, and databases such as SciFinder (www. cas.org/products/scifinder) provide information on the commercial availability of such compounds. Nevertheless, when applying these compounds, special consideration should be given to off-target effects, particularly given the difficulties in achieving specificity within phosphatase families. Of note, due to the evolutionary diversity of phosphatases, superfamilies like the PPPs, PTPs, and PPMs are usually not cross-inhibited when using small molecules but are sensitive to specific classes of inhibitors.9,157 In case of using a nonhydrolizable pSer/Thr peptide, cross-inhibition is possible since pSer/Thr activity is found within different superfamilies.6 The identification approaches and use of PTP inhibitors cover a broad chemical space ranging from small molecules to peptides and peptidomimetic compounds.134,164 Natural products and derivatives have served as a source for designing PTP inhibitors.164 Anionic pTyr-mimetic structures were successfully converted into PTP inhibitors: the most successful one is the difluorophosphonomethylphenylalanine (F2Pmp) group.168 In addition, two-site-binders as selective inhibitors of PTP1B have overcome the specificity problem emerging from J


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inhibit the PPP1R15A-PP1c and PPP1R15B-PP1c complexes, preventing the targeting of PP1 to eukaryotic initiation factor 2α, but it is still unclear if the molecule binds PP1c or the regulatory proteins.135,182 Subsequent structure activity relationship studies with this molecule led to a compound with enhanced cardioprotective activity that is 50-fold more potent than Salubrinal.183 Another work reported that the in vitro holoenzyme formation of PPP1R15A-PP1c was not disrupted through treatment with Guanabenz or Salubrinal, suggesting a mode of action in cells that the reconstruction in vitro did not fully mimic.184 As a molecular biology approach to selective PP1c inhibition, the PP1 inhibitory protein inhibitor 2 (PPP1R2) can be overexpressed or applied as recombinant protein for PP1c in vitro inhibition.185 A future direction of PPP inhibitors could follow the approach of targeting the oxidized state of PTPs:169 Recently, it was discovered that the metal ions in the active site of PP1c can be oxidized in cells through the reactive oxygen species-generating enzyme Nox4 rendering PP1 inactive.186 Designing ways to trigger this oxidation in a holoenzyme-specific manner for example by fusing Nox4 to regulatory proteins would open up a new approach to inhibit PP1 holoenzymes. For further information about the inhibition of PPPs, the reader is referred to reviews.11,155,157,160,187 Inhibitors of other phosphatase families are rare. One exception is the phosphatase Wip1 (PPM1D), which belongs to the PPM family and has been a major target due to its oncogenic activity.155 Recently, an orally active, allosteric inhibitor of Wip1 was introduced.188 Few phosphatases in the HAD phosphatase family have thus far been targeted. One example is Scp1, which is inhibited by rabeprazole that binds to a specific hydrophobic pocket on Scp1 and was identified by high throughput screening.189 Another example is the development of hydrolysis-resistant pyridoxal 5′-phosphate analogs as inhibitors of the pyridoxal phosphatase chronophin.190 The principle of nonhydrolizable substrate analogs and bioisosters for phosphatase inhibition has also been applied to PIP-phosphatases such as PTEN,131 for which also small molecule inhibitors were identified.156 Nevertheless, inhibitors of lipid phosphatases are still of very limited availability. Examples are inhibitors for inositol 5-phosphatases, including specific ones for OCRL/ INPP5B that were discovered by diverse screening approaches.191 Enzyme activators, or gain-of-function probes as equivalent to protein-overexpression but without increasing the protein levels, offer a unique way to address a particular cellular phenotype caused by this phosphatase, measuring its sufficiency to drive a particular cellular phenotype.158 The difficulty in designing activators is that one needs to know the exact mechanism of the enzyme, regarding for example an active conformation that can be stabilized or inhibiting the interaction with an inhibitory regulatory protein.43 Few activators are known, and most work has been done in this area on PPPs. An example for compounds leading to enhanced PP1c activity are the PP1-disrupting peptides (PDPs) that target a known regulatory site and disrupt interactions of PP1c with its regulatory proteins (i.e., the holoenzymes), releasing free PP1c that can dephosphorylate nearby substrates.160,192 Furthermore, the HDAC inhibitor (S)-8 acts by dissociating the cytosolic HDAC6-PP1c complex, allowing the release of PP1c that dephosphorylates protein kinase B (Akt), blocking downstream pro-survival signaling.185 Activators of PP2A that explore different mechanisms have been reported (reviewed in ref 187, also includes PP2A inhibitors). PP2A is inhibited by up-regulated proteins in cancer, and deinhibition or activation of PP2A is therefore

the high sequence homology of the catalytic site shared by the PTPs.165 Additional approaches include targeting the oxidized state of the catalytic cysteine by conformation-sensing antibodies (PTP1B),169 and allosteric sites on the catalytic domain or on other regions of the PTP. For example, allosteric inhibitors of PTP1B170 and SHP2171,172 lock these proteins in an inactive conformation, and these inhibitors were shown to be beneficial for cancer therapy. Another example of allosteric inhibition is a recent work on targeting the homotrimer complex of the DUSP PRL-1. The inhibitors disrupt this unique structural and regulatory property of PRL-1, which is essential for PRL-1-promoted cell proliferation and migration.173 The inhibition of PPPs has also been investigated intensively. A crucial difference to targeting PTPs is that not only the active site and allosteric sites on the PPP catalytic subunit (named after as “PP1c,” “PP2Ac,” and so on) but also the interacting proteins that form the holoenzymes with the catalytic subunits can be targeted. As is the case with PTPs, natural inhibitors of PPPs also posed as a foundation for designing active site inhibitors: okadaic acid, tautomycin, microcystins, nodularins, cantharidin, and fostriecin all served not only as a basis for rational inhibitor design11 but jointly as a common pharmacophore design which could make use of the advantages of each class.174 Of note, contrary to the PPPs PP1c, PP2Ac, PP4c, PP5c, and PP6c, due to unique regions in their catalytic domains, PP2Bc (also called PP3 or Calcineurin) and PP7c are insensitive to inhibition to such natural toxins.175 PP2Bc is instead selectively inhibited by FK506 and cyclosporine, which occupy a critical substrate recognition site but not the catalytically active site.176 Modeling, crystal structure information, and pharmacophore guided analog development has bettered inhibitor design for PPPs, thereby setting the core for a structure-based specificity hypothesis. The molecular docking approaches to the development of novel cantharidin analogues inhibiting PP1c, PP2Ac, and PP5c are an example.177 Newest developments in this field include structure−activity relationship studies of cantharidin analogs with PP1c, PP4c, and PP5c, including PP5c-inhibitor cocrystal structures, providing insights into a newly discovered selectivity of these compounds over PP4c.175 While the natural toxins are in general semiselective, fostriecin is a remarkable exception, being highly selective toward PP2Ac due to differences in the sequence of the proteins at fostriecin binding sites.178 The modest selectivity of most commercially available toxins, however, leads to the requirement of applying several inhibitors with different selectivity for the PPPs to be able to make a judgment on which PPP is involved in the dephosphorylation.157 As mentioned above, targeting the PPP holoenzymes is an alternative for PPP inhibition. The disruption of a PP1 holoenzyme by targeting PP1c was shown to inhibit its activity in HIV translation. The molecule binds presumably to the primary binding site of regulatory proteins to PP1c and selectively disrupts the association of PP1c and HIV-Tat protein in cells without affecting other PP1 holoenzymes tested in this context.179 Another new approach addressing the issue of specificity is targeting regulatory subunits of holoenzyme complexes, and it promises to enable achieving selectivity within the holoenzymes as shown for PP1 and the molecule Guanabenz.180 It was shown that the molecule Sephin1 (a more selective analog of Guanabenz) selectively binds to the stressinduced regulatory protein PPP1R15A, inhibiting its binding to PP1c and protecting cells from otherwise lethal protein misfolding stress.181 The molecule Salubrinal was reported to K


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ACS Chemical Biology desirable.3,159 Examples in this area are the compounds FTY720193 and its analog AAL(S), which have been shown to activate PP2A, and a recent study identified structural motifs required for cytotoxicity.194 These compounds were shown to be potent for treating acute myeloid leukemia cells193 and displayed synergistic effects when used in combination with tyrosine kinase inhibitors.195 Besides those for PPPs, other activators include the compound AQX-MN100 that selectively activates SHIP1, a PI(3,4,5)P3-phosphatase, by binding to an allosteric activation domain,196 and DEP-1 (PTPRJ), a receptor-like PTP, which is activated through induction of its dimerization by a peptide.197 Since many protein phosphatases are regulated by allosteric interactions with noncatalytic domains or autoinhibitory sites, targeting those sites holds promise for the successful design of new activators. To achieve complete specificity, Bishop et al. applied the “bump-hole” approach of allele-specific kinase inhibition to PTPs. They sensitized PTPs to biarsenical compounds creating via mutagenesis strategies non-natural allosteric-inhibition sites or changing the active sites, thereby controlling the function of each cellular phosphatase by designing selective mutant/ inhibitor pairs.198,199 Interestingly, recently they were able to apply that approach to create a SHP2 mutant that is activatable through binding of biarsenical compounds,200 showing the versatility of this approach.

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CONCLUSIONS AND OUTLOOK Many new methods that were either developed specifically for phosphatases or for a different area have entered the field of phosphatase research. While due to the complexity of these enzymes further methods still need to be developed to address specific issues, like the sensitivity of detecting phospholipid turnover, integrating the new methods reviewed here for phosphatase research has already and will continue to overcome the challenges attached to the research on phosphatases. This will help to substantially deepen our understanding of phosphatase biology by answering the key questions: (1) what are the substrates of a certain phosphatase; (2) what are the cellular roles of the phosphatase; and (3) what are the interaction partners, kinase counter players and networks of the phosphatase? This, in turn, combined with the development of new activators and inhibitors of phosphatases, promises to enable validating and using phosphatases as drug targets in the future.

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REFERENCES

(1) Cohen, P. T. W. (2004) Overview of protein serine/threonine phosphatases, in Protein Phosphatases (Ariño, J., and Alexander, D. R., Eds.), pp 1−20, Springer, Berlin. (2) Vintonyak, V. V., Waldmann, H., and Rauh, D. (2011) Using small molecules to target protein phosphatases. Bioorg. Med. Chem. 19, 2145−2155. (3) De Munter, S., Köhn, M., and Bollen, M. (2013) Challenges and opportunities in the development of protein phosphatase-directed therapeutics. ACS Chem. Biol. 8, 36−45. (4) Moorhead, G. B. G., De Wever, V., Templeton, G., and Kerk, D. (2009) Evolution of protein phosphatases in plants and animals. Biochem. J. 409, 401−409. (5) Manning, G., Plowman, G. D., Hunter, T., and Sudarsanam, S. (2002) Evolution of protein kinase signaling from yeast to man. Trends Biochem. Sci. 27, 514−520. (6) Li, X., Wilmanns, M., Thornton, J., and Köhn, M. (2013) Elucidating human phosphatase-substrate networks. Sci. Signaling 6, rs10. (7) Tonks, N. K. (2013) Protein tyrosine phosphatases - From housekeeping enzymes to master regulators of signal transduction. FEBS J. 280, 346−378. (8) Tonks, N. K. (2006) Protein tyrosine phosphatases: from genes, to function, to disease. Nat. Rev. Mol. Cell Biol. 7, 833−846.

ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acschembio.6b00570. Abbreviations, Glossary, and Supporting Table 1 (PDF)

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residues. These phosphatases are defined by the catalytic signature sequence Cx5R. PSTP: protein serine/threonine phosphatases, a large group of phosphatases, known to dephosphorylate phospho-serine and phospho-threonine residues. These phosphatases share a metal-dependent catalytic mechanism but are not evolutionary related between families within this group. DUSP: dual specificity phosphatases, a subfamily of the protein tyrosine phosphatases, which show diverse substrate specificity, dephosphorylating phospho-tyrosine containing proteins, phospho-serine/phospho-threonine, carbohydrates, mRNA, and phosphoinositides. HAD: haloacid dehalogenase phosphatases, characterized by a Rossmann-like fold and the active site sequence DxDx(V/T), contain enzymes with diverse substrate specificity such as toward pSer/Thr/Tyr, nucleotides, and pyridoxal-5′phosphate. Phosphoinositide phosphatases: phosphatases with activity toward phosphoinositides are found in different families and use different catalytic mechanisms. Specific tools to study them are included. Phosphatase substrates: approaches to identify protein and nonprotein substrates are discussed. These include massspectrometry-based methods like quantitative or targeted (phospho)proteomics, pull-down and coimmunoprecipitation methods, and proximity- and imaging-based approaches, most of which usually require the stabilization of the interaction. Phosphatase resources: computational resources for phosphatase research are given. Phosphatase functions: studying the functions of phosphatases using chemical modulators and sensors in cells is discussed. Chemical sensors: synthetic molecules used in biochemical assays to detect the presence or activity of a phosphatase, usually based on fluorescence. Chemical modulators: synthetic or natural molecules used as probes to either inhibit (loss-of-function probes) or enhance (gain-of-function probes) the activity of a phosphatase.

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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KEYWORDS PTP: protein tyrosine phosphatases, a large family of phosphatases named after the substrate specificity of many phosphatases in this family toward phospho-tyrosine L


Reviews

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Reviews

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Reviews

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Approaches to Study Phosphatases - ACS Publications - American

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