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Protein-specific imaging of posttranslational modifications
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ScienceDirect Protein-specific imaging of posttranslational modifications Wei Lin, Ling Gao and Xing Chen Protein posttranslational modifications (PTMs) modulate protein function, trafficking, and interactions. Many PTMs ubiquitously occurs on hundreds and thousands of proteins, which makes cellular imaging of the PTM state of a specific protein like looking for a needle in a haystack. A proximityenabled strategy, which exploits the spatial proximity between the PTM and the modified protein, has emerged as a valuable tool for protein-specific imaging of PTMs in single cells and tissue sections. The protein and the PTM are dually labeled with two distinct tags, which enable the generation of the nanometer proximity-dependent fluorescent signals for visualization. Herein, we review recent advances in the methodological developments and the applications of the proximity-enabled protein-specific imaging in studying phosphorylation, glycosylation, and lipidation. Address Beijing National Laboratory for Molecular Sciences, Key Laboratory of Bioorganic Chemistry and Molecular Engineering of Ministry of Education, College of Chemistry and Molecular Engineering, Synthetic and Functional Biomolecules Center, and Peking-Tsinghua Center for Life Sciences, Peking University, Beijing 100871, China Corresponding author: Chen, Xing ([email protected])
Current Opinion in Chemical Biology 2015, 28:156–163 This review comes from a themed issue on Synthetic biomolecules Edited by Christian Hackenberger and Peng Chen
http://dx.doi.org/10.1016/j.cbpa.2015.07.020 1367-5931/# 2015 Elsevier Ltd. All rights reserved.
Introduction The complexity of human proteome is dictated by posttranscriptional RNA processing and protein posttranslational modifications (PTMs) [1]. By alternative splicing, more than 80 000 mRNA transcripts are produced from approximately 20 000 human genes. After or concurrently with translation, more than 400 types of PTMs occur on proteins, which further amplifies the diversity of proteome by orders of magnitude [2]. PTMs are critical in regulating protein functions, localization, and interactions in various biological processes. It is the proteoforms (i.e. proteins bearing PTMs) that serve as the workhorses of a cell. Many important PTMs, such as phosphorylation, acetylation, glycosylation, and lipidation, involve Current Opinion in Chemical Biology 2015, 28:156–163
enzyme-catalyzed covalent addition of chemical groups onto amino acid side-chains. Quite often, a certain kind of PTM is ubiquitously found on hundreds and thousands of proteins in a mammalian cell. For example, it is estimated that about 30% of the human proteins are phosphorylated [3]; most of the cell-surface proteins are modified with glycans, which commonly share the same or similar structures [4]; and lipids are covalently bound to hundreds of proteins [5]. The past decade has witnessed an explosion in uncovering the proteome with a specific PTM, mainly facilitated by mass spectrometry-based proteomics [6,7]. For many PTMs, it is now feasible to perform large-scale analysis to identify tens of thousands of modification sites in cell or tissue samples. These largescale proteomic efforts have been generating PTM databases containing massive datasets, which lay the foundation for functional interpretations [8]. With the rapidly expanding PTM databases, a logical next step is to investigate the function of individual PTM sites and networks. To this end, one of the challenges is to develop methods for probing PTMs at the protein level to dissect how the biological function of a specific protein is regulated by its PTMs. Probing the PTM states of individual proteins has been commonly performed on purified proteins using biochemical assays, structural characterization, and mass spectrometry. However, these in vitro studies suffer from loosing the spatial information of proteins and are not well suited to address several important questions. What is the subcellular location for a protein PTM to occur? How is protein translocation regulated by PTMs? How do PTMs initiate and propagate cell signaling? To answer these questions, visualizing PTMs in a protein-specific manner in single cells is desired; however, the ubiquity of most PTMs makes this a highly challenging task. In this article, we review some of the recent methodology developments in protein-specific imaging of PTMs and the applications of these methods in studying the regulatory function of PTMs at the cellular level.
Protein-specific imaging of PTMs by the proximity-enabled strategy Technical principle and overview
The major technical challenge lies in how to selectively label or visualize the PTM of a specific protein in the presence of hundreds or thousands of other proteins carrying the same PTM. Due to the nongenetically encoded nature of most PTMs, the genetically encoded probes such as green fluorescent protein (GFP) cannot be tagged onto the PTM moieties. Alternatively, www.sciencedirect.com
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PTM-specific antibodies have been generated for some PTMs such as phosphorylation, acetylation, and methylation. Those PTM-specific antibodies are usually panspecific, recognizing PTMs on all proteins in a sequenceindependent manner. In some cases, it is possible to generate protein-specific and site-specific PTM antibodies, which can be applied for immunofluorescence imaging of PTMs in a protein-specific manner (also see examples described below). However, The specificity and affinity of many commercially available antibodies sometimes require careful validations [9], and it is troublesome and not always achievable to generate site-specific PTM antibodies for each protein. Furthermore, antibodies with good specificity and high affinity are rarely available for some PTMs including glycosylation and lipidation. A chemical reporter strategy has recently emerged for labeling and imaging PTMs that are difficult to target by antibodies [10,11]. In this strategy, the PTM substrates or building blocks, such as monosaccharides for glycosylation and lipids for lipidation, are chemically functionalized with a bioorthogonal chemical reporter (e.g. the azide or alkyne). The azide-modified or alkyne-modified PTM substrates or reporters are tolerated by the cellular enzymes and metabolically installed onto proteins, partially replacing their natural counterparts.
Subsequent chemical conjugation with a fluorophore by bioorthogonal reactions (e.g. click chemistry) enables visualization of PTMs in cells and animal tissues. However, like pan-specific antibodies, the chemical reporter strategy does not differentiate the identity of underlying proteins and therefore is not protein-specific for cellular imaging. Visualization of PTMs in a protein-specific manner would require integrating the identities of both the protein and the PTM. Built on the toolkits of PTM labeling and protein labeling, several dual labeling methods have been developed to achieve protein-specific imaging of phosphorylation [12,13,14], glycosylation [15,16, 17,18], and lipidation [19]. These methods are essentially based on a similar design principle, which exploits the spatial proximity between the PTM and the modified protein to endow pan-specific PTM labeling with protein specificity (Figure 1). The protein and PTM are labeled with two distinct tags, namely the protein tag and the PTM tag, respectively. The protein tag is specifically installed on the protein of interest by employing established protein labeling methods. In contrast, the PTM tag can be conjugated onto the PTM moieties attached on various proteins in a pan-specific manner.
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Proximity-enabled protein-specific imaging of PTMs. In the context of whole cell proteome, a PTM tag is installed, usually in a pan-specific manner, onto various proteins carrying this PTM, while the protein of interest is specifically labeled with a protein tag on the peptide scaffold. The protein and PTM tags on the protein of interest are in close proximity, which is exploited to in situ generate fluorescent signal through FRET or proximity ligation. The signal specificity is warranted by the stringent nanometer proximity requirement for effective FRET or proximity ligation. www.sciencedirect.com
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The PTM tag on the protein of interest then can be differentiated from the tags on other proteins by its close proximity with the protein tag. Two strategies, Fo¨rster resonance energy transfer (FRET) and proximity ligation, have been exploited to generate fluorescent signal confined only to the dually labeled protein. When the protein tag and the PTM tag are chosen to be two fluorophores that form a FRET pair, the intramolecular donor-acceptor distance fulfills the distance requirement for effective FRET (i.e. within 10 nm), whereas the intermolecular FRET is disfavored. Alternatively, the tags can be two nucleotides that are ligated together when bound in close proximity and the proximity ligation product is used to generate fluorescent signal for visualization.
proteome. In addition to pan-specific antibodies, sitespecific phosphorylation antibodies have also been produced for a variety of phosphorylated proteins [24]. There are now hundreds of phosphosite-specific antibodies that are commercially available. The applications of those antibodies include cellular immunofluorescence imaging of the phosphorylation state of specific proteins, such as histones [25], inner centromere protein [26], and adhesion-associated proteins [27]. Although the site-specific antibodies are a useful tool, the proximity-enabled protein-specific imaging strategies have been developed as a complementary means to visualize phosphorylation in cells and tissue sections. The Bastiaens group reported a proximity-enabled strategy based on FRET measured by FLIM (FLIM-FRET) for quantitative imaging of EGFR phosphorylation and signaling in live cells [12,13]. EGFR was genetically tagged with GFP and a Cy3-conjugated anti-phosphotyrosine antibody was microinjected into the cells expressing EGFR-GFP (Figure 2a). Efficient FRET only occurs when EGFR is phosphorylated and the antibody binding
Phosphorylation
Phosphorylation is probably the best studied PTM owing to its importance in cell signaling [20,21]. Using modern mass spectrometry-based proteomic analysis, large-scale profiling of thousands of unique phosphorylation sites can be performed in various mammalian cells [22,23], which underscores the ubiquity of phosphorylation in the Figure 2
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Protein-specific imaging of phosphorylation. (a) Schematic of dual labeling of EGFR for FLIM-FRET imaging of EGFR phosphorylation. The EGFRGFP expressing cells were labeled with a phosphotyrosine-specific Fab conjugated with Cy3. Efficient FRET only occurs between GFP and Cy3 bound on EGFR (i.e. intramolecular FRET). (b) A global analysis method was used to generate quantitative maps of the populations of EGFR-GFP in the phosphorylated and unphosphorylated states. The fluorescence lifetime of GFP in the unphosphorylated state, t1, is shortened to t2 through FRET when the EGFR-GFP is phosphorylated. The population maps (a1 and a2) are generated by analyzing the lifetimes in each pixel. (c) Populations of phosphorylated EGFR-GFP in live cells in response to the treatment of EGF-beads (red dots, top left panel). The color-coded scale indicates the population values of phosphorylated EGFR-GFP. The images are adapted with permission from Ref [12]. Current Opinion in Chemical Biology 2015, 28:156–163
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brings the Cy3 acceptor in close proximity to the GFP donor (i.e. within 10 nm range), which results in a shortening of the GFP fluorescence lifetime. Global analysis of FLIM images generates quantitative maps of the populations of EGFR in the phosphorylated and unphosphorylated states (Figure 2b). Using this strategy, the phosphorylation state of EGFR was monitored in live cells focally stimulated with epidermal growth factor (EGF) covalently attached to beads, which revealed lateral propagation of a signaling wave of EGFR phosphorylation and activation at the plasma membrane (Figure 2c) [12]. Together with theoretical analysis, quantitative imaging of EGFR phosphorylation was used to investigate the reaction network that couples EGFR activation to inhibition of protein tyrosine phosphatases (PTPs) [13]. Based on the imaging studies, it was proposed that EGF-induced H2O2 inhibits PTPs, which results in EGFR activation; diffusion of H2O2 in cells leads to lateral signal propagation. The FLIM-FRET methodology has also been applied to study phosphorylation of other proteins, including protein kinase Ca [28] and Fus3 (a component of mitogen-activated protein kinase (MAPK) signaling pathway in yeast) [29]. Furthermore, the Bastiaens group implemented an automated FLIM for sequential imaging of protein phosphorylation in cell assays, in which each spot contains cells expressing a distinct protein of interest tagged with GFP [30]. This high-throughput imaging setup allows in situ analysis of tyrosine phosphorylation networks and identification of components that relay signals from EGFR. Proximity ligation has also been exploited for generating the fluorescence signal for protein-specific imaging of phosphorylation [14]. The proximity ligation strategy was first developed by the Landegren group for sensitive detection of proteins [31]. A pair of proximity probes were produced by extending either the 50 or 30 end of a DNA apatamer recognizing platelet-derived growth factor Bchain (PDGF-BB) with two oligonucleotide extensions, which can be joined by ligation upon hybridization with a connector oligonucleotide. Proximal binding of the proximity probes to the homodimer of PDGF-BB promotes proximity ligation of the two extension oligonucleotides and the ligation product was detected with quantitative PCR. Other proteins are amenable to detection by proximity ligation given that a specific apatamer is available. The method was further extended by the Fredriksson to use antibody-oligonucleotide conjugates as the proximity probes, which largely expands the choices of the proteinrecognizing reagents [32]. An important technical advance in proximity ligation was later made by the Landegren group to visualize protein-protein interactions in individual cells, in which the two proximity probes were tailored to guide the formation of a circular DNA strand when bound in close proximity [33]. The DNA circle allows the use of localized rolling-circle amplification (RCA) to generate a randomly coiled, single-stranded DNA using one of the proximity probes as the primer. www.sciencedirect.com
The RCA product, which is therefore covalently linked to the target proteins, can be detected and visualized in situ by hybridization of complementary fluorescence-labeled oligonucleotides. To apply the RCA-based proximity ligation strategy for imaging the phosphorylation state of platelet-derived growth factor receptor b (PDGFRb), the So¨derberg group used two primary antibodies in cells and tissue sections to bind PDGFRb and the phosphorylated site, respectively, and the primary antibodies were then recognized by two secondary antibodies conjugated with the oligonucleotide pair [14]. Notably, a phosphoPDGFRb specific antibody was used in this work. A panspecific phosphorylation antibody could also be used in pair with the anti-PDGFRb antibody for imaging the phosphorylation of PDGFRb [34]. The methodology has been further applied to investigate the MARK2-dependent serine phosphorylation of Tau in NIH-3T3 cells [35], and the tyrosine phosphorylation of ABL, SHC, ERK2 and PI3K in response to kinase or phosphatase inhibitor treatment in K562 cells [36]. Glycosylation
Cell-surface proteins are mostly glycoproteins. N-linked and mucin-type O-linked glycosylation are two major types of glycosylation that occur in the endoplasmic reticulum (ER) and Golgi during the secretion of membrane and extracellular proteins [37,38]. In contrast to phosphorylation, cell-surface glycans in vertebrates are biopolymers constructed from nine kinds of monosaccharides that are connected via glycosidic linkages. Although N-linked and O-linked glycans can be further classified structurally into several subtypes, cell-surface proteins commonly carry glycans with the same or similar structures. Visualizing the glycosylation state of a specific protein of interest would greatly facilitate the studies on how glycosylation regulates protein functions in living cells. Several research groups have made important progresses toward this goal. The So¨derberg group adapted the in situ proximity ligation assay to image the sialyl-Tn antigen (Siaa2-6GalNAc-O-Ser/Thr) attached to the mucin protein MUC2 on tissue sections [15]. By using two antibodies against MUC2 and sialyl-Tn, respectively, the proximity ligation assay revealed that MUC2 is the major carrier of sialyl-Tn in intestinal metaplasia and gastric cancer. This methodology was further applied to detect various mucin glycoforms in mucinous adenocarcinomas by using a series of combinations of the available antibodies against mucin proteins including MUC1, MUC2, MUC5AC and MUC6, and against glycan antigens including Tn, Sialyl-Tn, T, Sialyl-Lea and Sialyl-LeX [39]. These studies demonstrate that protein-specific detection and imaging of glycans may potentially be valuable for cancer diagnosis, in complementary to using mucin proteins or O-glycan haptens alone as biomarkers. More recently, three research groups have reported the development of a FRET-based strategy, with some Current Opinion in Chemical Biology 2015, 28:156–163
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technical variations, for protein-specific imaging of glycosylation of cell-surface receptors in live cells [16,17,18]. The Suzuki group realized protein-specific imaging of sialylated glycans on EGFR and GLUT4 by metabolically labeling sialylated glycans with azides and genetically fusing GFP to the cytoplasmic terminal of proteins. Upon conjugation of the azides with tetramethylrhodamine (TMR)-alkyne, GFP and TMR formed a FRET donor-acceptor pair. Intramolecular FRET was detected and used to specifically image sialylated glycans on EGFR and GLUT4 [16]. However, this trans-membrane FRET strategy has two drawbacks. First, the FRET efficiency is relatively low since the donor is distant from the acceptor locating on the other side of the plasma membrane. Second, the acceptor spectral bleed-through may cause false positive signal in the FRET channel, since the acceptor is present in large excess. To overcome these problems, the Bertozzi group and our group independently developed a cismembrane FRET strategy. The Bertozzi group used the Fab fragments of antibodies that recognize the extracellular domains of cell-surface reporters to introduce the donor fluorophore, together with metabolic glycan labeling for introducing the acceptor fluorophore [17]. Furthermore, FLIM imaging was employed to circumvent the acceptor spectral bleed-through. The strategy was demonstrated by imaging sialylation of integrin avb3 in living cells and on tissue sections. Alternatively, our group developed a cis-membrane FRET-based strategy by exploiting a site-specific protein labeling method in conjugation with metabolic glycan labeling (Figure 3)
[18]. We adapted the PRIME protein labeling method, which uses an engineered lipoic acid ligase to recognize and functionalize a LAP peptide fused to the target protein with an azide-bearing lipoic acid derivative [40]. Two sequential click reactions were performed to install acceptor and donor onto glycans and the protein, respectively. The LAP peptide can be fused to the extracellular terminal of the target protein so that the donor and acceptor are both located on the same side of the membrane, ensuring close proximity for efficient FRET. To minimize the acceptor spectral bleed-through, we chose Fluor 488 and Alexa Fluor 647 as the donoracceptor pair, which possess well-separated excitation spectra. FRET imaging of the sialylation state of integrin aXb2 in live cells suggested that sialylation is important for aXb2 activation (Figure 3b). The generic nature of this imaging approach was further demonstrated by imaging glycosylation on additional cell-surface receptors including EGFR and TGF-b receptors. Lipidation
Protein lipidation modulates protein trafficking and localization and plays an important role in regulating underlying biological processes such as cell signaling [5]. Hundreds of eukaryotic proteins are covalently modified with lipids such as fatty acids and isoprenoids. For example, palmitoylome profiling using the acyl biotin exchange method [41,42] or the chemical reporter strategy [43,44] revealed several hundred palmitoylated protein in various cell types. Notably, lipidation-specific antibodies are generally difficult to generate and the lipid
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Protein-specific imaging of glycosylation. (a) Schematic of dual labeling of integrin aXb2 for cis-membrane FRET imaging of integrin sialylation. The protein is labeled with a FRET donor on its extracellular terminal by the PRIME protein labeling method, and the sialylated glycans are metabolically labeled with a FRET acceptor. To minimize the acceptor spectral bleed-through, Fluor 488 and Alexa Fluor 647 were chosen as the donor-acceptor pair, which possess well-separated excitation spectra. (b) FRET imaging of the sialylation state of integrain aXb2 in live cells while the activation state of aXb2 is probed by activation-dependent antibody MEM148. Scale bar: 5 mm. The images are adapted with permission from Ref [18]. Current Opinion in Chemical Biology 2015, 28:156–163
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chemical reporters provide a means to fluorescently label and visualize lipidation in cells [45,46]. Interestingly, the Fukata group recently developed an antibody specific for palmitoylated PSD-95, a landmark scaffold protein of the postsynaptic densities (PDs), and applied the antibody to visualize the palmitoylation state of PSD-95 in neurons [47]. Nevertheless, generation of lipid-specific antibodies remains challenging. Aiming to image Wnt palmitoylation, the Hannoush group developed a proximity ligation-based strategy for protein-specific imaging of palmitoylation (Figure 4) [19]. Mouse fibroblast L cells stably expressing Wnt3a were treated with 15-hexadecynoic acid (v-alkynyl palmitic acid, abbreviated as Alk-C16), an alkyne-functionalized palmitic acid analog, which was metabolically incorporated onto palmitoylated proteins. The alkyne was conjugated with a tag that is recognized by an antibody; Wnt3a protein was bound with a Wnt3a-specific antibody. Using two distinct secondary antibodies conjugated with complimentary oligonucleotides, the proximity ligation and RCA reaction were performed to generate a palmitoylated Wnt3a-specific fluorescent signal. The palmitoylation state of Wnt3a during secretion was visualized, which revealed that palmitoylated Wnt3a is
targeted to multivescular bodies (Figure 4b). The methodology has been generally applicable for other palmitoylated proteins such as sonic Hedgehog, tubulin, and H-Ras [48].
Conclusions and perspectives Protein-specific imaging of posttranslational modifications has been emerging as a useful tool to visualize the PTM state of a specific protein of interest in cells and in tissue sections. The two variations of the proximity-enabled strategy both rely on dual labeling of PTMs and proteins. To choose the imaging method, the advantages and limitations of each variant should be taken into consideration. First, the distance between the PTM tag and the protein tag is required to be within 10 nm and 40 nm for FRET and proximity ligation, respectively. More relaxed constraint on the proximity is generally prone to produce more non-specific fluorescence owing to the intermolecular signal. Nevertheless, extensive validation on the signal specificity should be performed for both methods. Second, the PTM level can be readily quantified by measuring the FRET fluorescence intensity. The ratio between the populations of modified and unmodified proteins can also be calculated in the FLIMFRET images. By contrast, the PTM level can only be
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Protein-specific imaging of lipidation. (a) Schematic of dual labeling of Wnt3a for proximity ligation and subsequent visualization of Wnt3a palmitoylation. The Wnt3a-expressing cells were metabolically labeled with Alk-C16, which was conjugated azide-OG488 that is recognized by an antibody; Wnt3a protein was bound with a Wnt3a-specific antibody. Using two distinct secondary antibodies conjugated with complimentary oligonucleotides, the proximity ligation and RCA reaction were performed to generate a palmitoylated Wnt3a-specific fluorescent signal. (b) Fluorescent imaging of the palmitoylation state of Wnt3a during secretion. CD 63 is an exosome marker, and CD9 and CD81 are MVB markers. Scale bar: 5 mm. The images are adapted with permission from Ref [19]. www.sciencedirect.com
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semi-quantitatively measured in the proximity ligationbased method by enumeration of the vesicle-like RCA products which represent the proximity amplification signal; the fluorescence intensity of the RCA products does not linearly depend on the PTM level. For the same reason, the ratio between modified and unmodified proteins cannot be measured, because the unmodified proteins are usually detected by immunofluorescence, a different readout from RCA. Third, the FRET-based method is compatible with live-cell imaging, while the proximity ligation-based variant is generally used for fixed cells due to the incompatibility of the PLA with live cells. Finally, although fluorescence microscopes with the FRET capability are now commonly accessible, the proximity ligation-based method only involves one fluorophore and hence demands less sophisticated instrumentation. In summary, the two variations of the proximity-enabled strategy should be chosen based on the specific systems under investigation and may be used in a complementary manner. We envision that the strategy will be further applied to protein-specific imaging of other types of PTMs, such as acetylation [49], methylation [50], and sulfenation [51]. Future methodology developments are sought to enable direct labeling of the PTM in a proteinspecific manner, which would largely improve the imaging processes.
Acknowledgements We acknowledge financial support from the National Natural Science Foundation of China (No. 91313301, No. 21425204, and No. 21172013), the National Basic Research Program of China (No. 2012CB917303), and the National Instrumentation Program (No. 2011YQ030124).
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43. Martin BR, Cravatt BF: Large-scale profiling of protein palmitoylation in mammalian cells. Nat Methods 2009, 6:135-138.
31. Fredriksson S, Gullberg M, Jarvius J, Olsson C, Pietras K, Gu´stafsdo´ttir SM, Ostman A, Landegren U: Protein detection using proximity-dependent DNA ligation assays. Nat Biotechnol 2002, 20:473-477. 32. Gullberg M, Gu´stafsdo´ttir SM, Schallmeiner E, Jarvius J, Bjarnega˚rd M, Betsholtz C, Landegren U, Fredriksson S: Cytokine detection by antibody-based proximity ligation. Proc Natl Acad Sci U S A 2004, 101:8420-8424. 33. So¨derberg O, Gullberg M, Jarvius M, Ridderstra˚le K, Leuchowius K-J, Jarvius J, Wester K, Hydbring P, Bahram F, Larsson L-G et al.: Direct observation of individual endogenous protein complexes in situ by proximity ligation. Nat Methods 2006, 3:995-1000. Integration of proximity ligation with RCA. 34. Leuchowius K-J, Jarvius M, Wickstro¨m M, Rickardson L, Landegren U, Larsson R, So¨derberg O, Frykna¨s M, Jarvius J: High content screening for inhibitors of protein interactions and post-translational modifications in primary cells by proximity ligation. Mol Cell Proteomics 2010, 9:178-183. 35. Gu GJ, Wu D, Lund H, Sunnemark D, Kvist AJ, Milner R, Eckersley S, Nilsson LNG, Agerman K, Landegren U et al.: Elevated MARK2-dependent phosphorylation of Tau in Alzheimer’s disease. J Alzheimers Dis 2013, 33:699-713. 36. Elfineh L, Classon C, Asplund A, Pettersson U, KamaliMoghaddam M, Lind SB: Tyrosine phosphorylation profiling via in situ proximity ligation assay. BMC Cancer 2014, 14:435.
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44. Yount JS, Moltedo B, Yang Y-Y, Charron G, Moran TM, Lo´pez CB, Hang HC: Palmitoylome profiling reveals S-palmitoylationdependent antiviral activity of IFITM3. Nat Chem Biol 2010, 6:610-614. 45. Hannoush RN, Sun J: The chemical toolbox for monitoring protein fatty acylation and prenylation. Nat Chem Biol 2010, 6:498-506. 46. Hang HC, Linder ME: Exploring protein lipidation with chemical biology. Chem Rev 2011, 111:6341-6358. 47. Fukata Y, Dimitrov A, Boncompain G, Vielemeyer O, Perez F, Fukata M: Local palmitoylation cycles define activity-regulated postsynaptic subdomains. J Cell Biol 2013, 202:145-161. 48. Gao X, Hannoush RN: Method for cellular imaging of palmitoylated proteins with clickable probes and proximity ligation applied to Hedgehog, tubulin, and Ras. J Am Chem Soc 2014, 136:4544-4550. Protein-specific imaging of lipidation in Hedgehog, tubulin, Ras. 49. Yang Y-Y, Ascano JM, Hang HC: Bioorthogonal chemical reporters for monitoring protein acetylation. J Am Chem Soc 2010, 132:3640-3641. 50. Islam K, Zheng W, Yu H, Deng H, Luo M: Expanding cofactor repertoire of protein lysine methyltransferase for substrate labeling. ACS Chem Biol 2011, 6:679-684. 51. Paulsen CE, Truong TH, Garcia FJ, Homann A, Gupta V, Leonard SE, Carroll KS: Peroxide-dependent sulfenylation of the EGFR catalytic site enhances kinase activity. Nat Chem Biol 2012, 8:57-64.
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ScienceDirect Protein-specific imaging of posttranslational modifications Wei Lin, Ling Gao and Xing Chen Protein posttranslational modifications (PTMs) modulate protein function, trafficking, and interactions. Many PTMs ubiquitously occurs on hundreds and thousands of proteins, which makes cellular imaging of the PTM state of a specific protein like looking for a needle in a haystack. A proximityenabled strategy, which exploits the spatial proximity between the PTM and the modified protein, has emerged as a valuable tool for protein-specific imaging of PTMs in single cells and tissue sections. The protein and the PTM are dually labeled with two distinct tags, which enable the generation of the nanometer proximity-dependent fluorescent signals for visualization. Herein, we review recent advances in the methodological developments and the applications of the proximity-enabled protein-specific imaging in studying phosphorylation, glycosylation, and lipidation. Address Beijing National Laboratory for Molecular Sciences, Key Laboratory of Bioorganic Chemistry and Molecular Engineering of Ministry of Education, College of Chemistry and Molecular Engineering, Synthetic and Functional Biomolecules Center, and Peking-Tsinghua Center for Life Sciences, Peking University, Beijing 100871, China Corresponding author: Chen, Xing ([email protected])
Current Opinion in Chemical Biology 2015, 28:156–163 This review comes from a themed issue on Synthetic biomolecules Edited by Christian Hackenberger and Peng Chen
http://dx.doi.org/10.1016/j.cbpa.2015.07.020 1367-5931/# 2015 Elsevier Ltd. All rights reserved.
Introduction The complexity of human proteome is dictated by posttranscriptional RNA processing and protein posttranslational modifications (PTMs) [1]. By alternative splicing, more than 80 000 mRNA transcripts are produced from approximately 20 000 human genes. After or concurrently with translation, more than 400 types of PTMs occur on proteins, which further amplifies the diversity of proteome by orders of magnitude [2]. PTMs are critical in regulating protein functions, localization, and interactions in various biological processes. It is the proteoforms (i.e. proteins bearing PTMs) that serve as the workhorses of a cell. Many important PTMs, such as phosphorylation, acetylation, glycosylation, and lipidation, involve Current Opinion in Chemical Biology 2015, 28:156–163
enzyme-catalyzed covalent addition of chemical groups onto amino acid side-chains. Quite often, a certain kind of PTM is ubiquitously found on hundreds and thousands of proteins in a mammalian cell. For example, it is estimated that about 30% of the human proteins are phosphorylated [3]; most of the cell-surface proteins are modified with glycans, which commonly share the same or similar structures [4]; and lipids are covalently bound to hundreds of proteins [5]. The past decade has witnessed an explosion in uncovering the proteome with a specific PTM, mainly facilitated by mass spectrometry-based proteomics [6,7]. For many PTMs, it is now feasible to perform large-scale analysis to identify tens of thousands of modification sites in cell or tissue samples. These largescale proteomic efforts have been generating PTM databases containing massive datasets, which lay the foundation for functional interpretations [8]. With the rapidly expanding PTM databases, a logical next step is to investigate the function of individual PTM sites and networks. To this end, one of the challenges is to develop methods for probing PTMs at the protein level to dissect how the biological function of a specific protein is regulated by its PTMs. Probing the PTM states of individual proteins has been commonly performed on purified proteins using biochemical assays, structural characterization, and mass spectrometry. However, these in vitro studies suffer from loosing the spatial information of proteins and are not well suited to address several important questions. What is the subcellular location for a protein PTM to occur? How is protein translocation regulated by PTMs? How do PTMs initiate and propagate cell signaling? To answer these questions, visualizing PTMs in a protein-specific manner in single cells is desired; however, the ubiquity of most PTMs makes this a highly challenging task. In this article, we review some of the recent methodology developments in protein-specific imaging of PTMs and the applications of these methods in studying the regulatory function of PTMs at the cellular level.
Protein-specific imaging of PTMs by the proximity-enabled strategy Technical principle and overview
The major technical challenge lies in how to selectively label or visualize the PTM of a specific protein in the presence of hundreds or thousands of other proteins carrying the same PTM. Due to the nongenetically encoded nature of most PTMs, the genetically encoded probes such as green fluorescent protein (GFP) cannot be tagged onto the PTM moieties. Alternatively, www.sciencedirect.com
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PTM-specific antibodies have been generated for some PTMs such as phosphorylation, acetylation, and methylation. Those PTM-specific antibodies are usually panspecific, recognizing PTMs on all proteins in a sequenceindependent manner. In some cases, it is possible to generate protein-specific and site-specific PTM antibodies, which can be applied for immunofluorescence imaging of PTMs in a protein-specific manner (also see examples described below). However, The specificity and affinity of many commercially available antibodies sometimes require careful validations [9], and it is troublesome and not always achievable to generate site-specific PTM antibodies for each protein. Furthermore, antibodies with good specificity and high affinity are rarely available for some PTMs including glycosylation and lipidation. A chemical reporter strategy has recently emerged for labeling and imaging PTMs that are difficult to target by antibodies [10,11]. In this strategy, the PTM substrates or building blocks, such as monosaccharides for glycosylation and lipids for lipidation, are chemically functionalized with a bioorthogonal chemical reporter (e.g. the azide or alkyne). The azide-modified or alkyne-modified PTM substrates or reporters are tolerated by the cellular enzymes and metabolically installed onto proteins, partially replacing their natural counterparts.
Subsequent chemical conjugation with a fluorophore by bioorthogonal reactions (e.g. click chemistry) enables visualization of PTMs in cells and animal tissues. However, like pan-specific antibodies, the chemical reporter strategy does not differentiate the identity of underlying proteins and therefore is not protein-specific for cellular imaging. Visualization of PTMs in a protein-specific manner would require integrating the identities of both the protein and the PTM. Built on the toolkits of PTM labeling and protein labeling, several dual labeling methods have been developed to achieve protein-specific imaging of phosphorylation [12,13,14], glycosylation [15,16, 17,18], and lipidation [19]. These methods are essentially based on a similar design principle, which exploits the spatial proximity between the PTM and the modified protein to endow pan-specific PTM labeling with protein specificity (Figure 1). The protein and PTM are labeled with two distinct tags, namely the protein tag and the PTM tag, respectively. The protein tag is specifically installed on the protein of interest by employing established protein labeling methods. In contrast, the PTM tag can be conjugated onto the PTM moieties attached on various proteins in a pan-specific manner.
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Proximity-enabled protein-specific imaging of PTMs. In the context of whole cell proteome, a PTM tag is installed, usually in a pan-specific manner, onto various proteins carrying this PTM, while the protein of interest is specifically labeled with a protein tag on the peptide scaffold. The protein and PTM tags on the protein of interest are in close proximity, which is exploited to in situ generate fluorescent signal through FRET or proximity ligation. The signal specificity is warranted by the stringent nanometer proximity requirement for effective FRET or proximity ligation. www.sciencedirect.com
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The PTM tag on the protein of interest then can be differentiated from the tags on other proteins by its close proximity with the protein tag. Two strategies, Fo¨rster resonance energy transfer (FRET) and proximity ligation, have been exploited to generate fluorescent signal confined only to the dually labeled protein. When the protein tag and the PTM tag are chosen to be two fluorophores that form a FRET pair, the intramolecular donor-acceptor distance fulfills the distance requirement for effective FRET (i.e. within 10 nm), whereas the intermolecular FRET is disfavored. Alternatively, the tags can be two nucleotides that are ligated together when bound in close proximity and the proximity ligation product is used to generate fluorescent signal for visualization.
proteome. In addition to pan-specific antibodies, sitespecific phosphorylation antibodies have also been produced for a variety of phosphorylated proteins [24]. There are now hundreds of phosphosite-specific antibodies that are commercially available. The applications of those antibodies include cellular immunofluorescence imaging of the phosphorylation state of specific proteins, such as histones [25], inner centromere protein [26], and adhesion-associated proteins [27]. Although the site-specific antibodies are a useful tool, the proximity-enabled protein-specific imaging strategies have been developed as a complementary means to visualize phosphorylation in cells and tissue sections. The Bastiaens group reported a proximity-enabled strategy based on FRET measured by FLIM (FLIM-FRET) for quantitative imaging of EGFR phosphorylation and signaling in live cells [12,13]. EGFR was genetically tagged with GFP and a Cy3-conjugated anti-phosphotyrosine antibody was microinjected into the cells expressing EGFR-GFP (Figure 2a). Efficient FRET only occurs when EGFR is phosphorylated and the antibody binding
Phosphorylation
Phosphorylation is probably the best studied PTM owing to its importance in cell signaling [20,21]. Using modern mass spectrometry-based proteomic analysis, large-scale profiling of thousands of unique phosphorylation sites can be performed in various mammalian cells [22,23], which underscores the ubiquity of phosphorylation in the Figure 2
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Protein-specific imaging of phosphorylation. (a) Schematic of dual labeling of EGFR for FLIM-FRET imaging of EGFR phosphorylation. The EGFRGFP expressing cells were labeled with a phosphotyrosine-specific Fab conjugated with Cy3. Efficient FRET only occurs between GFP and Cy3 bound on EGFR (i.e. intramolecular FRET). (b) A global analysis method was used to generate quantitative maps of the populations of EGFR-GFP in the phosphorylated and unphosphorylated states. The fluorescence lifetime of GFP in the unphosphorylated state, t1, is shortened to t2 through FRET when the EGFR-GFP is phosphorylated. The population maps (a1 and a2) are generated by analyzing the lifetimes in each pixel. (c) Populations of phosphorylated EGFR-GFP in live cells in response to the treatment of EGF-beads (red dots, top left panel). The color-coded scale indicates the population values of phosphorylated EGFR-GFP. The images are adapted with permission from Ref [12]. Current Opinion in Chemical Biology 2015, 28:156–163
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brings the Cy3 acceptor in close proximity to the GFP donor (i.e. within 10 nm range), which results in a shortening of the GFP fluorescence lifetime. Global analysis of FLIM images generates quantitative maps of the populations of EGFR in the phosphorylated and unphosphorylated states (Figure 2b). Using this strategy, the phosphorylation state of EGFR was monitored in live cells focally stimulated with epidermal growth factor (EGF) covalently attached to beads, which revealed lateral propagation of a signaling wave of EGFR phosphorylation and activation at the plasma membrane (Figure 2c) [12]. Together with theoretical analysis, quantitative imaging of EGFR phosphorylation was used to investigate the reaction network that couples EGFR activation to inhibition of protein tyrosine phosphatases (PTPs) [13]. Based on the imaging studies, it was proposed that EGF-induced H2O2 inhibits PTPs, which results in EGFR activation; diffusion of H2O2 in cells leads to lateral signal propagation. The FLIM-FRET methodology has also been applied to study phosphorylation of other proteins, including protein kinase Ca [28] and Fus3 (a component of mitogen-activated protein kinase (MAPK) signaling pathway in yeast) [29]. Furthermore, the Bastiaens group implemented an automated FLIM for sequential imaging of protein phosphorylation in cell assays, in which each spot contains cells expressing a distinct protein of interest tagged with GFP [30]. This high-throughput imaging setup allows in situ analysis of tyrosine phosphorylation networks and identification of components that relay signals from EGFR. Proximity ligation has also been exploited for generating the fluorescence signal for protein-specific imaging of phosphorylation [14]. The proximity ligation strategy was first developed by the Landegren group for sensitive detection of proteins [31]. A pair of proximity probes were produced by extending either the 50 or 30 end of a DNA apatamer recognizing platelet-derived growth factor Bchain (PDGF-BB) with two oligonucleotide extensions, which can be joined by ligation upon hybridization with a connector oligonucleotide. Proximal binding of the proximity probes to the homodimer of PDGF-BB promotes proximity ligation of the two extension oligonucleotides and the ligation product was detected with quantitative PCR. Other proteins are amenable to detection by proximity ligation given that a specific apatamer is available. The method was further extended by the Fredriksson to use antibody-oligonucleotide conjugates as the proximity probes, which largely expands the choices of the proteinrecognizing reagents [32]. An important technical advance in proximity ligation was later made by the Landegren group to visualize protein-protein interactions in individual cells, in which the two proximity probes were tailored to guide the formation of a circular DNA strand when bound in close proximity [33]. The DNA circle allows the use of localized rolling-circle amplification (RCA) to generate a randomly coiled, single-stranded DNA using one of the proximity probes as the primer. www.sciencedirect.com
The RCA product, which is therefore covalently linked to the target proteins, can be detected and visualized in situ by hybridization of complementary fluorescence-labeled oligonucleotides. To apply the RCA-based proximity ligation strategy for imaging the phosphorylation state of platelet-derived growth factor receptor b (PDGFRb), the So¨derberg group used two primary antibodies in cells and tissue sections to bind PDGFRb and the phosphorylated site, respectively, and the primary antibodies were then recognized by two secondary antibodies conjugated with the oligonucleotide pair [14]. Notably, a phosphoPDGFRb specific antibody was used in this work. A panspecific phosphorylation antibody could also be used in pair with the anti-PDGFRb antibody for imaging the phosphorylation of PDGFRb [34]. The methodology has been further applied to investigate the MARK2-dependent serine phosphorylation of Tau in NIH-3T3 cells [35], and the tyrosine phosphorylation of ABL, SHC, ERK2 and PI3K in response to kinase or phosphatase inhibitor treatment in K562 cells [36]. Glycosylation
Cell-surface proteins are mostly glycoproteins. N-linked and mucin-type O-linked glycosylation are two major types of glycosylation that occur in the endoplasmic reticulum (ER) and Golgi during the secretion of membrane and extracellular proteins [37,38]. In contrast to phosphorylation, cell-surface glycans in vertebrates are biopolymers constructed from nine kinds of monosaccharides that are connected via glycosidic linkages. Although N-linked and O-linked glycans can be further classified structurally into several subtypes, cell-surface proteins commonly carry glycans with the same or similar structures. Visualizing the glycosylation state of a specific protein of interest would greatly facilitate the studies on how glycosylation regulates protein functions in living cells. Several research groups have made important progresses toward this goal. The So¨derberg group adapted the in situ proximity ligation assay to image the sialyl-Tn antigen (Siaa2-6GalNAc-O-Ser/Thr) attached to the mucin protein MUC2 on tissue sections [15]. By using two antibodies against MUC2 and sialyl-Tn, respectively, the proximity ligation assay revealed that MUC2 is the major carrier of sialyl-Tn in intestinal metaplasia and gastric cancer. This methodology was further applied to detect various mucin glycoforms in mucinous adenocarcinomas by using a series of combinations of the available antibodies against mucin proteins including MUC1, MUC2, MUC5AC and MUC6, and against glycan antigens including Tn, Sialyl-Tn, T, Sialyl-Lea and Sialyl-LeX [39]. These studies demonstrate that protein-specific detection and imaging of glycans may potentially be valuable for cancer diagnosis, in complementary to using mucin proteins or O-glycan haptens alone as biomarkers. More recently, three research groups have reported the development of a FRET-based strategy, with some Current Opinion in Chemical Biology 2015, 28:156–163
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technical variations, for protein-specific imaging of glycosylation of cell-surface receptors in live cells [16,17,18]. The Suzuki group realized protein-specific imaging of sialylated glycans on EGFR and GLUT4 by metabolically labeling sialylated glycans with azides and genetically fusing GFP to the cytoplasmic terminal of proteins. Upon conjugation of the azides with tetramethylrhodamine (TMR)-alkyne, GFP and TMR formed a FRET donor-acceptor pair. Intramolecular FRET was detected and used to specifically image sialylated glycans on EGFR and GLUT4 [16]. However, this trans-membrane FRET strategy has two drawbacks. First, the FRET efficiency is relatively low since the donor is distant from the acceptor locating on the other side of the plasma membrane. Second, the acceptor spectral bleed-through may cause false positive signal in the FRET channel, since the acceptor is present in large excess. To overcome these problems, the Bertozzi group and our group independently developed a cismembrane FRET strategy. The Bertozzi group used the Fab fragments of antibodies that recognize the extracellular domains of cell-surface reporters to introduce the donor fluorophore, together with metabolic glycan labeling for introducing the acceptor fluorophore [17]. Furthermore, FLIM imaging was employed to circumvent the acceptor spectral bleed-through. The strategy was demonstrated by imaging sialylation of integrin avb3 in living cells and on tissue sections. Alternatively, our group developed a cis-membrane FRET-based strategy by exploiting a site-specific protein labeling method in conjugation with metabolic glycan labeling (Figure 3)
[18]. We adapted the PRIME protein labeling method, which uses an engineered lipoic acid ligase to recognize and functionalize a LAP peptide fused to the target protein with an azide-bearing lipoic acid derivative [40]. Two sequential click reactions were performed to install acceptor and donor onto glycans and the protein, respectively. The LAP peptide can be fused to the extracellular terminal of the target protein so that the donor and acceptor are both located on the same side of the membrane, ensuring close proximity for efficient FRET. To minimize the acceptor spectral bleed-through, we chose Fluor 488 and Alexa Fluor 647 as the donoracceptor pair, which possess well-separated excitation spectra. FRET imaging of the sialylation state of integrin aXb2 in live cells suggested that sialylation is important for aXb2 activation (Figure 3b). The generic nature of this imaging approach was further demonstrated by imaging glycosylation on additional cell-surface receptors including EGFR and TGF-b receptors. Lipidation
Protein lipidation modulates protein trafficking and localization and plays an important role in regulating underlying biological processes such as cell signaling [5]. Hundreds of eukaryotic proteins are covalently modified with lipids such as fatty acids and isoprenoids. For example, palmitoylome profiling using the acyl biotin exchange method [41,42] or the chemical reporter strategy [43,44] revealed several hundred palmitoylated protein in various cell types. Notably, lipidation-specific antibodies are generally difficult to generate and the lipid
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Protein-specific imaging of glycosylation. (a) Schematic of dual labeling of integrin aXb2 for cis-membrane FRET imaging of integrin sialylation. The protein is labeled with a FRET donor on its extracellular terminal by the PRIME protein labeling method, and the sialylated glycans are metabolically labeled with a FRET acceptor. To minimize the acceptor spectral bleed-through, Fluor 488 and Alexa Fluor 647 were chosen as the donor-acceptor pair, which possess well-separated excitation spectra. (b) FRET imaging of the sialylation state of integrain aXb2 in live cells while the activation state of aXb2 is probed by activation-dependent antibody MEM148. Scale bar: 5 mm. The images are adapted with permission from Ref [18]. Current Opinion in Chemical Biology 2015, 28:156–163
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chemical reporters provide a means to fluorescently label and visualize lipidation in cells [45,46]. Interestingly, the Fukata group recently developed an antibody specific for palmitoylated PSD-95, a landmark scaffold protein of the postsynaptic densities (PDs), and applied the antibody to visualize the palmitoylation state of PSD-95 in neurons [47]. Nevertheless, generation of lipid-specific antibodies remains challenging. Aiming to image Wnt palmitoylation, the Hannoush group developed a proximity ligation-based strategy for protein-specific imaging of palmitoylation (Figure 4) [19]. Mouse fibroblast L cells stably expressing Wnt3a were treated with 15-hexadecynoic acid (v-alkynyl palmitic acid, abbreviated as Alk-C16), an alkyne-functionalized palmitic acid analog, which was metabolically incorporated onto palmitoylated proteins. The alkyne was conjugated with a tag that is recognized by an antibody; Wnt3a protein was bound with a Wnt3a-specific antibody. Using two distinct secondary antibodies conjugated with complimentary oligonucleotides, the proximity ligation and RCA reaction were performed to generate a palmitoylated Wnt3a-specific fluorescent signal. The palmitoylation state of Wnt3a during secretion was visualized, which revealed that palmitoylated Wnt3a is
targeted to multivescular bodies (Figure 4b). The methodology has been generally applicable for other palmitoylated proteins such as sonic Hedgehog, tubulin, and H-Ras [48].
Conclusions and perspectives Protein-specific imaging of posttranslational modifications has been emerging as a useful tool to visualize the PTM state of a specific protein of interest in cells and in tissue sections. The two variations of the proximity-enabled strategy both rely on dual labeling of PTMs and proteins. To choose the imaging method, the advantages and limitations of each variant should be taken into consideration. First, the distance between the PTM tag and the protein tag is required to be within 10 nm and 40 nm for FRET and proximity ligation, respectively. More relaxed constraint on the proximity is generally prone to produce more non-specific fluorescence owing to the intermolecular signal. Nevertheless, extensive validation on the signal specificity should be performed for both methods. Second, the PTM level can be readily quantified by measuring the FRET fluorescence intensity. The ratio between the populations of modified and unmodified proteins can also be calculated in the FLIMFRET images. By contrast, the PTM level can only be
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Protein-specific imaging of lipidation. (a) Schematic of dual labeling of Wnt3a for proximity ligation and subsequent visualization of Wnt3a palmitoylation. The Wnt3a-expressing cells were metabolically labeled with Alk-C16, which was conjugated azide-OG488 that is recognized by an antibody; Wnt3a protein was bound with a Wnt3a-specific antibody. Using two distinct secondary antibodies conjugated with complimentary oligonucleotides, the proximity ligation and RCA reaction were performed to generate a palmitoylated Wnt3a-specific fluorescent signal. (b) Fluorescent imaging of the palmitoylation state of Wnt3a during secretion. CD 63 is an exosome marker, and CD9 and CD81 are MVB markers. Scale bar: 5 mm. The images are adapted with permission from Ref [19]. www.sciencedirect.com
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semi-quantitatively measured in the proximity ligationbased method by enumeration of the vesicle-like RCA products which represent the proximity amplification signal; the fluorescence intensity of the RCA products does not linearly depend on the PTM level. For the same reason, the ratio between modified and unmodified proteins cannot be measured, because the unmodified proteins are usually detected by immunofluorescence, a different readout from RCA. Third, the FRET-based method is compatible with live-cell imaging, while the proximity ligation-based variant is generally used for fixed cells due to the incompatibility of the PLA with live cells. Finally, although fluorescence microscopes with the FRET capability are now commonly accessible, the proximity ligation-based method only involves one fluorophore and hence demands less sophisticated instrumentation. In summary, the two variations of the proximity-enabled strategy should be chosen based on the specific systems under investigation and may be used in a complementary manner. We envision that the strategy will be further applied to protein-specific imaging of other types of PTMs, such as acetylation [49], methylation [50], and sulfenation [51]. Future methodology developments are sought to enable direct labeling of the PTM in a proteinspecific manner, which would largely improve the imaging processes.
Acknowledgements We acknowledge financial support from the National Natural Science Foundation of China (No. 91313301, No. 21425204, and No. 21172013), the National Basic Research Program of China (No. 2012CB917303), and the National Instrumentation Program (No. 2011YQ030124).
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Current Opinion in Chemical Biology 2015, 28:156–163