Proteomic identification of protein tyrosine phosphatase and substrate

Subscriber access provided by UNIV OF LOUISIANA

Article

Proteomic identification of protein tyrosine phosphatase and substrate interactions in living mammalian cells by genetic encoding of irreversible enzyme inhibitors Hongting Tang, Zhen Dai, Xuewen Qin, Wenkang Cai, Liming Hu, Yujia Huang, Wenbing Cao, Fan Yang, Chu Wang, and Tao Liu J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.8b06922 • Publication Date (Web): 24 Sep 2018 Downloaded from http://pubs.acs.org on September 24, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 8 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society

Proteomic identification of protein tyrosine phosphatase and substrate interactions in living mammalian cells by genetic encoding of irreversible enzyme inhibitors Hongting Tang1,†, Zhen Dai1,2,†, Xuewen Qin1, Wenkang Cai1, Liming Hu1, Yujia Huang1, Wenbing Cao1,2, Fan Yang3, Chu Wang3, and Tao Liu1,* 1

State Key Laboratory of Natural and Biomimetic Drugs, School of Pharmaceutical Sciences, Peking University, 38 Xueyuan Road, Haidian District, Beijing 100191, China 2 Department of Chemistry, State Key Laboratory of Elemento-Organic Chemistry, Nankai University, Tianjin 300071, China 3 College of Chemistry and Molecular Engineering, Peking University, Beijing, 100871, China *Correspondence to: [email protected] Supporting Information ABSTRACT: Protein tyrosine phosphatases (PTPs) play critical roles in cell signaling pathways, but identification of unknown PTPs for a given substrate in live cells remain technically challenging. Here, we synthesized a series of tyrosine-based irreversible PTP inhibitors and characterized by site-specific encoding on substrate proteins in cells with an expanded genetic code. By finetuning the chemical reactivity, we identified optimal active amino acid probes to covalently cross-link a PTP and its substrate both in vitro and in mammalian cells. Using HER2 as an example, we provide first direct evidence of HER2 Y1023 and SHP2 crosslinking in situ in living human cells. Moreover, proteomic analysis using our approach identified PTP1B as a novel phosphatase for HER2 that specifically dephosphorylated pY1221 position, which may shed light on the puzzle of PTP1B’s role in HER2 positive breast cancer. This novel method provides a useful tool for dissecting tyrosine phosphor-regulation in living cells.

INTRODUCTION Protein tyrosine phosphorylation is a common posttranslational modification that plays a pivotal role in modulating a variety of cellular processes1-2. The reversible nature of tyrosine phosphorylation is tightly regulated by the opposing activities of protein tyrosine kinases (PTKs) and protein tyrosine phosphatases (PTPs), and not surprisingly, the malfunction of this regulation results in a number of human diseases. The regulation of PTK and substrate interactions in disease conditions have been well documented, and over two dozen drugs that target PTKs are in clinical use today3. In contrast, even though the human genome encodes 103 PTPs4, a number comparable to the number of human PTKs (90)5, the function and regulation of the human PTP-substrate network remain largely unexplored for the following reasons: (1) the importance and specificity of PTP-substrate interactions in signaling and disease have historically been unappreciated6; (2) studying such interactions is difficult because detection of phosphate group removal requires the phosphorylation event to have happened first7-8; (3) the interactions are highly transient7, 9; and (4) most importantly, PTP-substrate interactions in cells depend not only on intrinsic enzymatic specificity but also on their subcellular co-localization and protein-protein interactions in situ10. Therefore, although extensive studies have been performed in vitro to elucidate the substrate specificities of PTPs11-12, the lack of suitable methods for directly detecting PTP-substrate pairs in live cells has markedly ham-

pered the identification of novel interactions and the discovery of potential therapeutic PTP targets9, 13. One of the most extensively used approaches for studying PTPs in a cellular context is the substrate-trapping mutant approach, which is used to capture binding complexes between catalytically inactive PTPs and their substrates14. However, this approach relies on the extent of substrate tyrosine phosphorylation and provides only indirect evidence for potential interactions for multiprotein complexes. More importantly, the approach can be used only to identify the substrates of a given PTP, whereas researchers are more interested in determining the PTPs responsible for the phosphorregulations on a protein of interest involved in development or disease conditions. To better understand the functions and regulations of phosphotyrosine (pY) signaling, new methods for identifying the PTPs responsible for a given substrate in living cells are urgently needed. In our laboratory, we are interested in HER2 (human epidermal growth factor receptor 2) protein, an EGFR family member involved in the progression of various cancers, most prominently breast cancer. Like other EGFR family members, HER2 has multiple tyrosine residues on its intracellular region that are controlled by reversible phosphorylation (Figure 1a) and that mediate diverse cell signaling by binding to various adaptor proteins15. Research suggests that the phosphorylation statuses of these residues are regulated by different PTPs16-18 and most of them are yet to be determined. To this end, we

ACS Paragon Plus Environment

Journal of the American Chemical Society 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

have developed a method using mechanism-based noncanonical amino acid cross-linking coupled with mass spectrometry, for identifying novel PTPs for a given substrate in living human cells. PTPs share a reaction mechanism that involves nucleophilic catalysis by an active-site cysteine residue with a low pKa (Figure 1b). Many covalent PTP inhibitors with reactive groups capable of forming covalent adducts with the thiolate group of the active-site cysteine have been developed, and these work have been extensively reviewed19. Recently, advances in the development of an expanded genetic code approach have allowed several reactive noncanonical amino acids (ncAAs) to be genetically encoded into proteins in bacterial or mammalian cell culture by using cognate orthogonal amber suppressor tRNA and aminoacyl-tRNA synthetase (RS) pairs20-24. We hypothesized that genetic encoding of irreversible cysteine reactive PTP inhibitors as warheads at specific phosphorylation sites on substrate proteins would trap the PTP enzymes by formation of covalent enzyme-substrate complexes in situ in the proteomes of mammalian cells, therefore be very useful in identifying unknown PTPs for a substrate of interest by proteomic analysis. RESULTS AND DISCUSSION Design, synthesis, and activity analysis of ncAAs that react with PTPs. On the basis of existing irreversible inhibitors of PTPs19, we designed and synthesized six tyrosine analogues, each containing one of five types of cysteine-reactive groups (Figure 1c). Specifically, ClpAcF, its less-reactive derivative FpAcF, and the nonreactive control compound pAcF were based on α-haloacetophenone-type PTP inhibitors25-26 (Figure S1 and S2). AcrF was selected as a Michael acceptor type PTP cross-linker27-28. BetY is a haloalkane-type crosslinker that readily react with cysteine. Because alkynes have been shown to act as irreversible inhibitors for enzymes containing active-site cysteine 29-30, the alkyne containing OPrY and the less-reactive alkene containing OAlY were selected as potential cross-linkers for PTPs. Finally, OMeY was included as a nonreactive and nonphosphorylatable control. First, we tested the inhibitory activities of the designed ncAAs against PTP. As free amino acids, ClpAcF, FpAcF, AcrF, and BetY efficiently inhibited SHP2-PTP activity, a model PTP used for in vitro study, whereas OPrY and OAlY had no observable effects (Figure 2a). To determine whether the inhibitory activity resulted from covalent cross-linking, we used electrospray ionization mass spectrometry (ESI-MS) to obtain the exact mass of SHP2-PTP in the presence and absence of the ncAAs. FpAcF was selected as a model compound as it is readily to be incorporated in mammalian cell culture22. As shown in Figure 2b, incubation with FpAcF resulted in an extra peak at 46283.6 Da (a mass shift of +205 Da relative to the peak observed in the absence of FpAcF), corresponding to a FpAcF-cysteine reaction adduct. In comparison, the mass spectrum after incubation with pAcF, a nonreactive analogue of FpAcF, with SHP2-PTP under the same conditions gave only a single peak (at 46078.6 Da), corresponding to the unmodified protein (Figure S3). The SHP2 catalytic domain contains eight cysteines, so to confirm that the reaction occurred at the active-site cysteine (C459), we allowed a C459A mutant (SHP2CA) to react with FpAcF under the same conditions, and no cross-linking adduct was observed (Figure S4). These results indicate that FpAcF inhibited the PTP by forming a covalent adduct specific to the active-site cysteine

and thus might be useful as a cross-linking probe to capture PTP-substrate interactions.

Figure 1. Overview of the strategy to cross-link PTPs and HER2 in live cells. (a) Scheme showing multiple phosphorylation sites on HER2 intracellular domain and in situ cross-linking strategy for capturing potential interacting PTPs in mammalian cells. (b) Reaction mechanism of PTP catalytic cysteine cross-linking with the ncAA probes. (c) Structures of ncAAs used in this study.

In vitro cross-linking of recombinant substrate proteins containing ncAAs. Next, we determined whether purified substrate proteins containing our ncAAs in place of pYs could cross-link with PTPs in vitro. The genetic incorporation these ncAAs in E. coli were first examined with a previously developed polyspecific Methanococcus jannaschii TyrRS (MjPolyRS)/tRNACUA pair31. Nonsense codon suppression was evaluated by means of recombinant expression of superfolder green fluorescent protein (sfGFP) mutants bearing a permissive amber codon at D134, which is surface-exposed to cytosolic nucleophiles (Figure S5). In the presence of 1 mM ncAAs, substantial fluorescence enhancement was observed for all the tested compounds (Figure 2c).


Page 2 of 8

Page 3 of 8 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society

Figure 2. Characterization of ncAA probes as free amino acids and on recombinant proteins. (a) PTP inhibition assay. SHP2 catalytic domain at 25 µM concentration was incubated with different concentrations of ncAAs and measured for activities using pnitrophenylphoshpate (pNPP) as a substrate. Data represent mean of three independent experiments ± s.d. (b) ESI-MS analysis of SHP2-PTP (calculated mass 46079.2 Da, observed mass 46078.6 Da) treated with 1 mM FpAcF. An observed peak of 46283.6 Da matches with a SHP2-FpAcF reaction adduct (calculated mass 46284.2 Da). (c) Evaluating the nonsense suppression efficiencies with different ncAAs in E coli. using a polyspecific MjPolyRS/tRNACUA pair. Fluorescence intensities of GFP mutants in the crude cell lystate were recorded using a fluorescent plate reader. (d) ESI-MS analysis of GFP mutant containing FpAcF (calculated mass 27989.6 Da, observed mass 27989.9 Da). The observed peak of 27858.5 Da represents the GFP mutant without a Nterminal methionine (calculated mass 27858.4 Da), which is typically seen for recombinant protein produced in E coli. The observed peak of 28277.6 Da matches a covalent attachment of a glutathione molecule (calculated mass 28276.6 Da).

The GFP mutants were then individually purified, and their yields are shown in Table S1. ESI-MS was performed to confirm the incorporation of the ncAAs in the sfGFP mutants. The observed masses were consistent with the calculated masses for all the ncAAs except AcrF and ClpAcF, which were too reactive to stay intact in the cytosol after genetic encoding (Figure S6) and were therefore excluded from further experiments. It is worth mentioning that in the spectrum of sfGFPD134FpAcF, we observed an extra peak (28277.6 Da) that was consistent with the formation of glutathione adducts after nucleophilic reaction with the fluorine atom (Figure 2d). Although part of the FpAcF-containing protein was modified, we used it, along with BetY, OPrY, and OAlY, for further experiments. Next, we attempted to recombinantly express PTP substrate proteins containing the remaining ncAAs. Although we are interested in HER2, recombinant expression of its intracellular domain turned out to be a challenge. Therefore, we chose the SH3 domain of Abelson murine leukemia viral oncogene homolog 1 (ABL1) that we previously characterized as a recombinant model protein containing pY32. ABL1-SH3 Y70 mutants bearing different ncAAs were expressed in E. coli and verified by ESI-MS (Figure S7). Cross-linking experiments

were performed in vitro by incubating ABL1-SH3 mutants with SHP2-PTP. SDS-PAGE analysis of the ABL1 mutants containing BetY and FpAcF under denaturing conditions showed an extra band with a molecular weight corresponding to the SHP2-ABL complex. Replacing FpAcF with pAcF, which differs by one fluorine atom, eliminated the band. In addition, no band was observed upon incubation of ABL1Y70FpAcF with the active-site mutant SHP2CA under the same conditions (Figure 3a). These results confirm crosslinking between our ncAAs and the active-site cysteine of the selected PTP. Because the extra band was much weaker than we expected, we determined the binding affinity between the two proteins by means of biolayer interferometry, using the nonfluorinated substrate ABL1-Y70pAcF (Figure S8). This experiment confirmed that the binding was indeed very weak (measured affinity, 37 uM), which explains the intensity of the band observed on the SDS-PAGE gel. This is expected as only the catalytic domain was used for in vitro binding measurement, where adaptor domains, such as SH2 and SH3 domains were often involved for interactions in live cells. It is worth mentioning that affinity measurement using fluorinated ABL1-Y70FpAcF with SHP2 resulted in an unmeasurable Koff value, which is consistent with covalent binding.

Figure 3. Covalent cross-linking of PTP and substrate in vitro and in live cells. (a) SDS-PAGE analysis of complexes formation by incubating SHP2 with ABL1 mutants containing different ncAAs. ABL1 mutants containing BetY or FpAcF at position 70 formed covalent bands with SHP2, indicated by red arrow. SHP2CA stands for active-site mutation of Cys to Ala. (b) Genetic incorporation of different ncAAs in EGFP-Y39TAG expressed in mammalian cells. (c) Construction of plasmids encoding SHP2 and HER2 Y1023TAG mutant. SHP2 expression vector contains a RS/tRNA expression cassette, which consists a CMV promoterdriven EcpAcFRS and eight copies of H1 promoter-driven BstRNACUA. (d) Western blot analysis of SHP2-HER2 complexes formation in live cells. V5-tagged SHP2 and FLAG-tagged HER2 mutants were co-expressed in HEK293 cells. The complex was immunoprecipitated with an anti-FLAG antibody, and visualized using an anti-V5 antibody. The red arrows indicate the crosslinking band.

PTP-substrate cross-linking in mammalian cells. Next, we sought to determine whether substrate proteins containing



the ncAAs could cross-link PTPs in the proteomes of live mammalian cells. We began by genetically incorporating these ncAAs in HEK293 cells using an evolved E. coli pAcFRS/tRNACUA pair33-34, which has been shown to accept many structurally similar tyrosine analogues as substrates in mammalian cells35. A plasmid encoding one copy of CMV promoter-driven EcpAcFRS and eight copies of an H1 promoter-driven Bacillus stearothermophilus tRNACUA 34 (BstRNAPyl) cassette was co-transfected with a reporter plasmid pEGFP-Y39TAG (Figure S9). Bright green fluorescence was observed under a microscope in the presence of all the tested ncAAs, suggesting the production of full-length EGFP by amber suppression (Figure 3b). With this tool in hand, we moved on to investigate the cross-linking of PTPsubstrate pairs in cells. It has been reported that dephosphorylation of the pY1023 residue of HER2 is controlled by SHP216. Therefore, we generated a HER2 mutant construct by mutating the codon of Y1023 to the amber stop codon (Figure 3c). The mutant proteins containing the various ncAAs were co-expressed with CMV-driven full-length SHP2 in HEK293 cells, and the crosslinking band was analyzed by a western blot assay. As shown in Figure 3d, a clear cross-linking band, with a size consistent with the molecular weight of the SHP2-HER2 complex, was observed for both the HER2-Y1023FpAcF and the HER2Y1023BetY mutants. In contrast, no extra band was detected for the HER2-Y1023 mutants containing OMeY, OAIY, and OPrY (Figure S10), suggesting that the reactivity of the alkyne and alkene functionalities was too weak for PTP cross-linking. Surprisingly, we found that the covalent band generated by the BetY-containing mutant was more intense than that generated by the FpAcF-containing mutant. This result may be due to glutathione addition to FpAcF prior to triggering of the dephosphorylation event as we observed for recombinant proteins (Figure S11). These results highlight the importance of fine-tuning the chemical reactivity of the ncAA towards PTPs under different conditions. Identification of PTP1B as a potential HER2 partner by cross-linking coupled with proteomic analysis. We wondered whether our cross-linking strategy could be used in combination with proteomic analysis to explore unknown PTP-substrate interactions in live mammalian cells. We are interested in HER2 Y1221/Y1222 residues, whose phosphorylation statuses are important for HER2 signaling and are correlated to poor survival rates in breast cancer patients36-39. Currently, there is no confirmed PTP responsible for the dephosphorylation of these two sites, and both their functions and regulations cannot be distinguished. To identify the PTPs responsible for these sites, we first constructed an expression plasmid containing a CMV promoter-driven FLAG-tagged HER2-Y1221TAG mutant and the entire RS/tRNA cassette (Figure 4). HER2-Y1221 mutants were overexpressed in HEK293 cells in the presence of 250 uM BetY or OMeY. Proteins associated with the HER2 mutants were co-immunoprecipitated, separated by SDS-PAGE under denaturing conditions, and analyzed by silver staining. Extra bands with molecular weights larger than that of HER2 were observed for BetY containing mutant (Figure 4), indicating potential cross-linked adducts. Proteomic analysis of these bands gave a total of 358 and 310 proteins for each mutant respectively. Investigation of the 116 unique proteins associated with the HER2-Y1221BetY mutant identified one PTP enzyme, PTP1B (tyrosine-protein phosphatase non-

receptor type 1, encoded by PTN1). In addition, the top 50 proteins based on ranking score were analyzed for other potential covalent adducts (Table S2). Enzymes with activesite cysteines, such as protein disulfide isomerase (PDI) family members (PDIA6, PDIA4, PDIA3, and PDIA1), were among the top hits on the list. PDIs are enzymes found in the eukaryotic endoplasmic reticulum, where they catalyze the formation of disulfide bonds to facilitate correct folding of proteins. Likewise, the fuctions of the other proteins at the top of the list suggest that they may be involved in HER2 translation, folding, transportation, glycosylation, degradation and other processes. These proteins might have been trapped by the BetY residue when they interacted with the HER2. The rest of the proteins may be rich in surface-exposed cysteine residues, located in close vicinity to HER2, or present in cells at high abundances, and therefore underwent random crosslinking to the HER2 mutant.

Figure 4. Mechanism-based ncAA cross-linking coupled with proteomic analysis for HER2 Y1221 position in situ in live cells. CMV promoter-driven HER2 mutants containing either BetY or OMeY residues at 1221 position were expressed in HEK293 cells. Proteins with a molecular weight larger than HER2 were visualized on a silver staining gel and isolated for proteomic analysis.

Site-specific interaction of PTP1B with HER2 at Y1221. The proteomics analysis results are exciting because to the best of our knowledge, HER2 is not known to be a substrate for PTP1B. Although numerous studies have suggested an important role of PTP1B in HER2 positive breast cancer40-41, there is no evidence that PTP1B directly dephosphorylated HER2 in cells. To confirm our results, we co-expressed PTP1B and HER2-Y1221BetY in HEK293 cells, purified the covalent adduct by immunoprecipitation, and analyzed it by a western blot assay. As shown in Figure 5a and Figure S12, PTP1B was covalently attached to the HER2-Y1221 mutant containing BetY, whereas no cross-linked band was observed for the control mutant containing OMeY. In addition, the C215S mutant of PTPB1 fails to covalently cross-link with the unnatural mutant HER2 under the same condition, which rule out the possibility that PTPB1 could be cross-linking through another (non-active-site) residue to HER2 (Figure S13). The results were further validated using three substrate-trapping mutants of PTP1B (D181A42, D181A/Y46A43, and C215S44). As shown in Figure 5b, the assay revealed that more HER2 protein was pulled down by the trapping mutants than by wildtype PTP1B, suggesting complex formation in cells. To further


Page 4 of 8

Page 5 of 8 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society prove PTP1B is responsible for the dephosphorylation of HER2 pY1221, HER2 were co-expressed with PTP1B or inactive mutant PTP1B-C215S. Western blot analysis showed a decrease of phosphorylation signal for wild-type PTP1B as compared to the inactive mutant (Figure S14), suggesting that PTP1B indeed dephosphorylates HER2 on Y1221and therefore validating our results.

Figure 5. Verification of PTP1B and HER2-1221 interaction in cells. (a) Western blot analysis of PTP1B co-expression with HER2 mutants containing either BetY or OMeY at 1221 position. (b) PTP1B substrate-trapping mutants were co-expressed with FLAG-tagged HER2 in HEK293 cells. The complexes were immunoprecipitated with anti-PTP1B antibody and immunoblotted by anti-Flag antibody. (c) Western blot analysis of PTP1B coexpression with HER2 mutants containing BetY at different phosphorylation sites. A strong covalent band can only be visualized with HER2 Y1221BetY mutant. (d) Crystal structure of the PTP1B binding to a substrate peptide derived from insulin receptor containing tandem phosphotyrosines (PDB code 1G1F).

Because there are multiple phosphorylation sites (Y877, Y1023, Y1139, Y1196, Y1221, Y1222 and Y1248) on the tail of the intracellular domain of HER2, we were curious to know whether PTP1B could interact with other nearby pY residues. HER2 mutant plasmids containing a pY to ncAA mutation were constructed by replacing the tyrosine codon with the amber codon. The mutants, along with Y1221, were coexpressed with PTP1B in the presence of BetY. Interestingly, western blot assay showed that none of these mutants gave a strong cross-linking band, except Y1221 (Figures 5c and S15).

Analysis of known PTP1B substrates revealed a subgroup containing tandem pYs, including the insulin receptor Y1150/1151 and tropomyosin receptor kinase B Y706/70745-46. Close examination of the PTP1B structure bound to an insulin receptor substrate peptide containing tandem pYs suggested that PTP1B possesses a positive charged binding pocket in addition to the catalytic site47. As shown in Figure 5d, PTP1B binds strongly to the second pY residue via salt bridges between the phosphate group and the side chains of Arg24 and Arg254. Such interactions help to position the first pY residue (or the BetY residue in our application) at the active site of PTP1B for a cysteine-mediated reaction to take place. This speculation was further supported with HER2 variant containing a Y1222F mutation (Figure S16). In conclusion, PTP1B interacts with HER2 in live cells site-specifically at the 1221 residue. CONCLUSIONS Identifying the PTPs responsible for phosphorylating a given substrate in human cells is challenging. To overcome this challenge, we have developed a cross-linking strategy for directly capturing the cysteine-tyrosine interactions between PTP active sites and substrates in situ. The key to this advance is genetic encoding of cysteine-reactive ncAAs sitespecifically into substrate proteins in human cells to covalently trap the enzymes on the basis of their common reaction mechanism. Because the reactivities of the ncAAs must be finetuned within a cellular context, we screened a panel of six tyrosine analogues containing one of five types of cysteinereactive groups: haloacetophenone (FpAcF and ClpAcF), Michael acceptor (AcrF), haloalkane (BetY), alkyne (OPrY), and alkene (OAlY). Unfortunately, ClpAcF and AcrF, which were based on existing PTP inhibitors, were too reactive to be retained on the cytosolic portion of the substrate protein. FpAcF, a less-reactive haloacetophenone-type PTP inhibitor, showed the best cross-linking reactivity with PTPs in vitro. Interestingly, in human cell culture, the haloalkane-type ncAA BetY, which has relatively low reactivity, showed cross-linking results superior to those of FpAcF, suggesting that the electrophilic ncAA needed to be fine-tuned within a cellular environment. By covalently cross-linking the active-site cysteine with the pY-mimetic probe, our method enables direct observation of the formation of PTP-substrate complexes in situ in living mammalian cells, thus providing unambiguous evidence for assigning PTP-substrate pairs. To the best of our knowledge, such assignments cannot be made by any of the existing methods used to study PTPs. Existing methods, such as the substrate-trapping mutant method, rely heavily on the extent of substrate phosphorylation and capture entire complexes, thus, identifying the right substrate is difficult if multiple proteins are involved. Other methods, such as the use of smallmolecule chemical cross-linkers, cannot capture the highly transient interactions between PTPs and substrates in cells. For a proof-of-concept study, we chose the HER2-Y1023 and SHP2 pair and used western blotting to directly visualize in situ formation of the covalent SHP2-HER2 complex for the first time. Finally, by coupling in-cell cross-linking with proteomic analysis of the HER2-Y1221 residue, we identified PTP1B as a novel phosphatase partner, a result that may shed light on the puzzle of the role of PTP1B in HER2-positive breast cancer. Interestingly, this previously unreported interac-



tion was very specific to the Y1221 residue, which validates the utility of our approach for identifying PTPs for individual tyrosine residues on a substrate of interest. In summary, we have developed a new method to study PTPs in mammalian cells using an expanded genetic code. By covalently cross-linking the active-site cysteine with the pYmimetic probe, the method captures the transient PTPsubstrate interactions into stable complexes in live cells, thus providing unambiguous evidence for assigning PTP-substrate pairs. This method does not require a kinase to phosphorylate the substrate in advance and works in situ in complex proteomes of living mammalian cells, thus will be useful for revealing the human PTP-substrate network and improving our understanding of the phosphor-regulation of important tyrosine residues.

ASSOCIATED CONTENT Supporting Information Experimental details and supporting figures. This material is available free of charge via the Internet at http://pubs.acs.org.”

AUTHOR INFORMATION Corresponding Author [email protected] ORCID: Tao Liu: 0000-0001-5347-5892

Author Contributions †These authors contributed equally.

ACKNOWLEDGMENT This work was financially supported by National Key Research and Development Program of China (No. 2016YFA0201400), the National Natural Science Foundation of China (21778005), Peking University Health Science Center (BMU20160537 and BMU2017QQ006). T.L. thanks the Youth Thousand-Talents Program of China for support. The authors acknowledge Dr. Wen Zhou at the mass spectrometry facility of National Center for Protein Sciences at Peking University for the assistance in proteomic experiments, and Jun Li at the State Key Laboratory of Natural and Biomimetic Drugs for assistance in high resolution protein mass spectrometry.

REFERENCES (1) Pawson, T.; Scott, J. D., Protein phosphorylation in signaling-50 years and counting. Trends Biochem Sci 2005, 30 (6), 286-90. (2) Hunter, T., Tyrosine phosphorylation: thirty years and counting. Curr Opin Cell Biol 2009, 21 (2), 140-6. (3) Wu, P.; Nielsen, T. E.; Clausen, M. H., Small-molecule kinase inhibitors: an analysis of FDA-approved drugs. Drug Discov Today 2016, 21 (1), 5-10. (4) Alonso, A.; Sasin, J.; Bottini, N.; Friedberg, I.; Friedberg, I.; Osterman, A.; Godzik, A.; Hunter, T.; Dixon, J.; Mustelin, T., Protein tyrosine phosphatases in the human genome. Cell 2004, 117 (6), 699711. (5) Robinson, D. R.; Wu, Y. M.; Lin, S. F., The protein tyrosine kinase family of the human genome. Oncogene 2000, 19 (49), 55485557. (6) Yu, Z. H.; Zhang, Z. Y., Regulatory Mechanisms and Novel Therapeutic Targeting Strategies for Protein Tyrosine Phosphatases. Chem Rev 2018, 118 (3), 1069-1091. (7) De Munter, S.; Kohn, M.; Bollen, M., Challenges and opportunities in the development of protein phosphatase-directed therapeutics. ACS Chem Biol 2013, 8 (1), 36-45.

(8) Tonks, N. K., Protein tyrosine phosphatases--from housekeeping enzymes to master regulators of signal transduction. FEBS J 2013, 280 (2), 346-78. (9) Fahs, S.; Lujan, P.; Kohn, M., Approaches to Study Phosphatases. ACS Chem Biol 2016, 11 (11), 2944-2961. (10) Hale, A. J.; Ter Steege, E.; den Hertog, J., Recent advances in understanding the role of protein-tyrosine phosphatases in development and disease. Dev Biol 2017, 428 (2), 283-292. (11) Ren, L.; Chen, X.; Luechapanichkul, R.; Selner, N. G.; Meyer, T. M.; Wavreille, A. S.; Chan, R.; Iorio, C.; Zhou, X.; Neel, B. G.; Pei, D., Substrate specificity of protein tyrosine phosphatases 1B, RPTPalpha, SHP-1, and SHP-2. Biochemistry 2011, 50 (12), 2339-56. (12) Zhang, Z. Y., Protein tyrosine phosphatases: structure and function, substrate specificity, and inhibitor development. Annu Rev Pharmacol Toxicol 2002, 42, 209-34. (13) Tiganis, T.; Bennett, A. M., Protein tyrosine phosphatase function: the substrate perspective. Biochem J 2007, 402 (1), 1-15. (14) Flint, A. J.; Tiganis, T.; Barford, D.; Tonks, N. K., Development of "substrate-trapping" mutants to identify physiological substrates of protein tyrosine phosphatases. Proc Natl Acad Sci U S A 1997, 94 (5), 1680-5. (15) Roskoski, R., Jr., The ErbB/HER family of protein-tyrosine kinases and cancer. Pharmacol Res 2014, 79, 34-74. (16) Zhou, X.; Agazie, Y. M., Molecular mechanism for SHP2 in promoting HER2-induced signaling and transformation. J Biol Chem 2009, 284 (18), 12226-34. (17) Wang, H. M.; Xu, Y. F.; Ning, S. L.; Yang, D. X.; Li, Y.; Du, Y. J.; Yang, F.; Zhang, Y.; Liang, N.; Yao, W.; Zhang, L. L.; Gu, L. C.; Gao, C. J.; Pang, Q.; Chen, Y. X.; Xiao, K. H.; Ma, R.; Yu, X.; Sun, J. P., The catalytic region and PEST domain of PTPN18 distinctly regulate the HER2 phosphorylation and ubiquitination barcodes. Cell Res 2014, 24 (9), 1067-90. (18) Li, H.; Yang, F.; Liu, C.; Xiao, P.; Xu, Y.; Liang, Z.; Liu, C.; Wang, H.; Wang, W.; Zheng, W.; Zhang, W.; Ma, X.; He, D.; Song, X.; Cui, F.; Xu, Z.; Yi, F.; Sun, J. P.; Yu, X., Crystal Structure and Substrate Specificity of PTPN12. Cell Rep 2016, 15 (6), 1345-58. (19) Ruddraraju, K. V.; Zhang, Z. Y., Covalent inhibition of protein tyrosine phosphatases. Mol Biosyst 2017, 13 (7), 1257-1279. (20) Xiang, Z.; Lacey, V. K.; Ren, H. Y.; Xu, J.; Burban, D. J.; Jennings, P. A.; Wang, L., Proximity-Enabled Protein Crosslinking through Genetically Encoding Haloalkane Unnatural Amino Acids. Angew Chem Int Edit 2014, 53 (8), 2190-2193. (21) Furman, J. L.; Kang, M. C.; Choi, S.; Cao, Y.; Wold, E. D.; Sun, S. B.; Smider, V. V.; Schultz, P. G.; Kim, C. H., A Genetically Encoded aza-Michael Acceptor for Covalent Cross-Linking of Protein-Receptor Complexes. J Am Chem Soc 2014, 136 (23), 84118417. (22) Xiang, Z.; Ren, H. Y.; Hu, Y. S.; Coin, I.; Wei, J.; Cang, H.; Wang, L., Adding an unnatural covalent bond to proteins through proximity-enhanced bioreactivity. Nat Methods 2013, 10 (9), 885-+. (23) Li, F.; Zhang, H.; Sun, Y.; Pan, Y.; Zhou, J.; Wang, J., Expanding the genetic code for photoclick chemistry in E. coli, mammalian cells, and A. thaliana. Angew Chem Int Ed Engl 2013, 52 (37), 9700-4. (24) Lee, Y. J.; Wu, B.; Raymond, J. E.; Zeng, Y.; Fang, X.; Wooley, K. L.; Liu, W. R., A genetically encoded acrylamide functionality. ACS Chem Biol 2013, 8 (8), 1664-70. (25) Arabaci, G.; Guo, X. C.; Beebe, K. D.; Coggeshall, K. M.; Pei, D., alpha-Haloacetophenone derivatives as photoreversible covalent inhibitors of protein tyrosine phosphatases. J Am Chem Soc 1999, 121 (21), 5085-5086. (26) Arabaci, G.; Yi, T.; Fu, H.; Porter, M. E.; Beebe, K. D.; Pei, D. H., alpha-Bromoacetophenone derivatives as neutral protein tyrosine phosphatase inhibitors: Structure-activity relationship. Bioorg Med Chem Lett 2002, 12 (21), 3047-3050. (27) Seiner, D. R.; LaButti, J. N.; Gates, K. S., Kinetics and mechanism of protein tyrosine phosphatase 1B inactivation by acrolein. Chem Res Toxicol 2007, 20 (9), 1315-20. (28) Liu, S. J.; Zhou, B.; Yang, H. Y.; He, Y. T.; Jiang, Z. X.; Kumar, S.; Wu, L.; Zhang, Z. Y., Aryl vinyl Sulfonates and Sulfones as


Page 6 of 8

Page 7 of 8 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society active site-directed and mechanism-based probes for protein tyrosine phosphatases. J Am Chem Soc 2008, 130 (26), 8251-8260. (29) Ekkebus, R.; van Kasteren, S. I.; Kulathu, Y.; Scholten, A.; Berlin, I.; Geurink, P. P.; de Jong, A.; Goerdayal, S.; Neefjes, J.; Heck, A. J. R.; Komander, D.; Ovaa, H., On Terminal Alkynes That Can React with Active-Site Cysteine Nucleophiles in Proteases. J Am Chem Soc 2013, 135 (8), 2867-2870. (30) Sommer, S.; Weikart, N. D.; Linne, U.; Mootz, H. D., Covalent inhibition of SUMO and ubiquitin-specific cysteine proteases by an in situ thiol-alkyne addition. Bioorgan Med Chem 2013, 21 (9), 2511-2517. (31) Young, D. D.; Young, T. S.; Jahnz, M.; Ahmad, I.; Spraggon, G.; Schultz, P. G., An evolved aminoacyl-tRNA synthetase with atypical polysubstrate specificity. Biochemistry 2011, 50 (11), 1894-900. (32) Luo, X.; Fu, G.; Wang, R. E.; Zhu, X.; Zambaldo, C.; Liu, R.; Liu, T.; Lyu, X.; Du, J.; Xuan, W.; Yao, A.; Reed, S. A.; Kang, M.; Zhang, Y.; Guo, H.; Huang, C.; Yang, P. Y.; Wilson, I. A.; Schultz, P. G.; Wang, F., Genetically encoding phosphotyrosine and its nonhydrolyzable analog in bacteria. Nat Chem Biol 2017, 13 (8), 845-849. (33) Chin, J. W.; Cropp, T. A.; Anderson, J. C.; Mukherji, M.; Zhang, Z.; Schultz, P. G., An expanded eukaryotic genetic code. Science 2003, 301 (5635), 964-7. (34) Liu, W. S.; Brock, A.; Chen, S.; Chen, S. B.; Schultz, P. G., Genetic incorporation of unnatural amino acids into proteins in mammalian cells. Nat Methods 2007, 4 (3), 239-244. (35) Chatterjee, A.; Xiao, H.; Bollong, M.; Ai, H. W.; Schultz, P. G., Efficient viral delivery system for unnatural amino acid mutagenesis in mammalian cells. Proc Natl Acad Sci U S A 2013, 110 (29), 11803-8. (36) Lucs, A. V.; Muller, W. J.; Muthuswamy, S. K., Shc is required for ErbB2-induced inhibition of apoptosis but is dispensable for cell proliferation and disruption of cell polarity. Oncogene 2010, 29 (2), 174-87. (37) Marone, R.; Hess, D.; Dankort, D.; Muller, W. J.; Hynes, N. E.; Badache, A., Memo mediates ErbB2-driven cell motility. Nat Cell Biol 2004, 6 (6), 515-22. (38) Jones, R. B.; Gordus, A.; Krall, J. A.; MacBeath, G., A quantitative protein interaction network for the ErbB receptors using protein microarrays. Nature 2006, 439 (7073), 168-74. (39) Frogne, T.; Laenkholm, A. V.; Lyng, M. B.; Henriksen, K. L.; Lykkesfeldt, A. E., Determination of HER2 phosphorylation at tyro-

sine 1221/1222 improves prediction of poor survival for breast cancer patients with hormone receptor-positive tumors. Breast Cancer Res 2009, 11 (1). (40) Julien, S. G.; Dube, N.; Read, M.; Penney, J.; Paquet, M.; Han, Y. X.; Kennedy, B. P.; Muller, W. J.; Tremblay, M. L., Protein tyrosine phosphatase 1B deficiency or inhibition delays ErbB2-induced mammary tumorigenesis and protects from lung metastasis. Nat Genet 2007, 39 (3), 338-346. (41) Bentires-Alj, M.; Neel, B. G., Protein-tyrosine phosphatase 1B is required for HER2/Neu-induced breast cancer. Cancer Res 2007, 67 (6), 2420-2424. (42) Lu, X. Q.; Malumbres, R.; Shields, B.; Jiang, X. Y.; Sarosiek, K. A.; Natkunam, Y.; Tiganis, T.; Lossos, I. S., PTP1B is a negative regulator of interleukin 4-induced STAT6 signaling. Blood 2008, 112 (10), 4098-4108. (43) Boubekeur, S.; Boute, N.; Pagesy, P.; Zilberfarb, V.; Christeff, N.; Issad, T., A New Highly Efficient Substrate-trapping Mutant of Protein Tyrosine Phosphatase 1B (PTP1B) Reveals Full Autoactivation of the Insulin Receptor Precursor. Journal of Biological Chemistry 2011, 286 (22), 19373-19380. (44) Zhang, Y. L.; Yao, Z. J.; Sarmiento, M.; Wu, L.; Burke, T. R.; Zhang, Z. Y., Thermodynamic study of ligand binding to proteintyrosine phosphatase 1B and its substrate-trapping mutants. Journal of Biological Chemistry 2000, 275 (44), 34205-34212. (45) Bandyopadhyay, D.; Kusari, A.; Kenner, K. A.; Liu, F.; Chernoff, J.; Gustafson, T. A.; Kusari, J., Protein-tyrosine phosphatase 1B complexes with the insulin receptor in vivo and is tyrosinephosphorylated in the presence of insulin. Journal of Biological Chemistry 1997, 272 (3), 1639-45. (46) Ozek, C.; Kanoski, S. E.; Zhang, Z. Y.; Grill, H. J.; Bence, K. K., Protein-tyrosine Phosphatase 1B (PTP1B) Is a Novel Regulator of Central Brain-derived Neurotrophic Factor and Tropomyosin Receptor Kinase B (TrkB) Signaling. Journal of Biological Chemistry 2014, 289 (46), 31682-92. (47) Salmeen, A.; Andersen, J. N.; Myers, M. P.; Tonks, N. K.; Barford, D., Molecular basis for the dephosphorylation of the activation segment of the insulin receptor by protein tyrosine phosphatase 1B. Mol Cell 2000, 6 (6), 1401-12.



Page 8 of 8

SYNOPSIS TOC


8

Proteomic identification of protein tyrosine phosphatase and substrate

Recommend Documents