Cytocompatible Modification of Thermoresponsive ... - ACS Publications

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Cytocompatible Modification of Thermoresponsive Polymers on Living Cells for Membrane Proteomic Isolation and Analysis Yuanzi Wu, Shuigen Wu, Shanyun Ma, Fen Yan, and Zuquan Weng Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b04201 • Publication Date (Web): 11 Feb 2019 Downloaded from http://pubs.acs.org on February 11, 2019

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Analytical Chemistry

Cytocompatible Modification of Thermoresponsive Polymers on Living Cells for Membrane Proteomic Isolation and Analysis Yuanzi Wu*, Shuigen Wu†, Shanyun Ma†, Fen Yan, and Zuquan Weng* College of Biological Science and Engineering, Fuzhou University, Fuzhou, Fujian 350002, China. ABSTRACT: Efficient strategies for enriching and separating proteins are important and challenging for membrane proteomics. Many existing methods are caught in the dilemma of preserving maximal membrane proteins while avoiding the contamination of cytoplasmic proteins and organelles. Here, we report a polymer anchoring strategy for the selective preparation of membrane proteins through cell surface-initiated atom transfer radical polymerization. The cytocompatible polymerization strategy enables thermoresponsive p(NIPAAm) chains to be grown from a specific protein on the surface of living cells. The polymer tagged membrane protein could be easily separated and enriched by thermoprecipitation. This method led to the identification of 1825 proteins of which 1036 (71.7%) were specific membrane proteins in the E. coli. The separated proteins were identified by 2-DE and mass spectrometry. Among the 12 proteins spots from the gel slice, eight were identified as outer membrane proteins. The described strategy opens up a new avenue for membrane protein enriching and separating, and may expedite the future development of membrane proteomics. Membrane proteins play important roles in many fundamental biological processes, such as material transport, signal transduction, and cell-cell communication.

1, 2

Membrane proteins have been targeted for drug design since they account for a large proportion of therapeutic drug targets.3 Identification of membrane proteins in cells would provide insight into disease mechanisms and potential therapeutic targets. Currently, mass spectrometry-based proteomics is playing a central role in elucidating protein components, structures, and functional relationships.4, 5 Although this strategy is exceptionally powerful now, the membrane proteome still faces analytical challenges

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owing to low abundance, insolubility, and heterogeneity. For a focused analysis of cell membrane subproteomes, prior enrichment and purification of the membrane fraction are desired. Various strategies have been developed for enriching membrane proteins, however, they are still far from being perfect. For example, density gradient centrifugation used to separate the membrane fraction always carries over subcellular organelles, resulting in interference in the proteomic analysis.6 To improve the coverage as well as the accuracy of membrane proteins, a number of improved isolation methods have been reported,3 and these can be briefly divided into two categories, namely, physisorption and chemical labeling. Physisorption of materials, such as colloidal silica nanoparticles7-9 and magnetic nanoparticles,10, 11 onto the cell membrane is driven by an electrostatic force. This simple method ensures efficient membrane fraction capture only under moderate washing conditions without detergents.12 Moreover, some nanoparticles may also be internalized in cells and contaminated by cytosolic components. Covalent coupling is based on the covalent binding of cell surface proteins with desired molecules. This method involves biotinylating the amino groups of membrane proteins and subsequently affinity isolation by using streptavidin beads 13-15 or agarose resin16-17. Fang et al. specifically isolated glycosylated membrane proteins by covalently binding oxidized glycan moieties to hydrazide-functionalized microspheres.18 However, some challenges remain, such as part of the small tag molecules might pass through the cell membrane, leading to the capture of intracellular proteins. Compared with modification by small molecules, engineering cell surfaces with macromolecules has proved to be a more promising approach for endowing cells with new functions and characteristics.19-22 Functional polymers were traditionally introduced onto the cell membrane by a “grafting-to” approach.23-26 However, this strategy is limited due to the low grafting efficiency of the macromolecules onto the cell surface.25 The “grafting from” strategy offers an improved polymer brush, which could be initiated and grown from the cell membrane directly.20, 27, 28 This approach can increase the polymer density and control the chain length while maintaining cell


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functions. Moreover, though part of the initiators might pass across the membrane and react with the cytosolic components, this interference could be neglected since the polymerization mainly initiated outside the cell membrane. Here, we demonstrate a method for cytocompatible polymer modification on the cell surface and apply it to enrich for proteomic analysis of cell membrane proteins. This approach involves the covalent immobilization of an initiator on cell membrane proteins, in situ polymerization from viable cells, and the thermoprecipitation purification of membrane proteins described in our previous work (scheme 1).29 During this process, the proteome of interest is covalently tagged with the thermoresponsive polymer, while the membranes remain intact. The enriched proteins were identified by SDS-PAGE and mass spectrometry with a high membrane protein yield of 71.7%. The results indicated that this novel strategy might be of promise in cell membrane proteomic research. EXPERIMENTAL SECTION Materials and Reagents N,N,N',N'-tetramethylethylenediamine,

bis-acrylamide,

iodoacetamide,

and

ammonium persulfate were purchased from Sigma. N-Isopropyl acrylamide (NIPPAm) was purchased from TCI. Tris(2-chloroethyl) phosphate (TCEP) and N,N,N',N'',N''pentamethyldiethylenetriamine (PMDETA) was purchased from Aladdin. Nsuccinimidyl-3-(2-pyridyldithiol)propionate (SPDP) was purchased from Huateng (China). DL-Dithiothreitol, ascorbic acid, glycine, acrylamide, thiourea, ethylene diamine tetraacetic acid, and tris (hydroxymethyl) aminomethane were purchased from Sangon Biotech (Shanghai). Yeast extract and tryptone were purchased from Oxoid. All the other chemical reagents were supplied by Sinopharm (China). Preparation of E. coli protoplast E.coli BL21 cells from a single colony on an agar plate were first inoculated in 5



mL of autoclaved yeast extract peptone LB liquid media. After incubating for 14 h at 37 °C, the E.coli cells were collected by centrifugation at 873 ×g for 10 min at 4 °C. The collected pellet was washed twice with a 10.3% sucrose solution, then suspended in 5 mL of P buffer with 5 mg/mL of lysozyme at 37 °C for up to 3 h. The pellet was further centrifuged and washed in P buffer three times to remove the supernatant. The protoplast was finally suspended in P buffer at a concentration of ~ 200 mg/mL before further usage. Initiator anchoring and cell surface-initiated polymerization 0.5 mL of SPDP (70 mM, DMSO) was added into a 10 mL protoplast suspension dropwise, then mixed by stirring rapidly. After reacting for 10 min, the mixture was centrifuged at 388 ×g for 5 min. The supernatant was removed by a pipette. The procedure was repeated with P buffer three times. 10 mL of TCEP (60 mM, P buffer) was then mixed and reacted with the SPDP-modified protoplast solution in the same manner. The pellet was resuspended in 10 mL of P buffer after the centrifugation and washing steps. Bis[2-(2’-bromoisobutyryloxy)ethyl]disulfide (500 μL, 0.12 mM) was added dropwise into the TCEP treated protoplast solution, then mixed by stirring gently. After reacting for 30 min, the mixture was centrifuged and washed. The pellet was collected and stored in 10 mL of P buffer as the final initiator modified protoplast. The polymerization solution was prepared in a sealed flask by mixing 10 mL water, 5 mL MeOH, ~6

mg PMDETA, ~6 mg CuCl2, and 1 g NIPPAm,

(PMDETA/CuCl2/NIPPAm = 1/1/255). After deoxygenation with nitrogen for approximately 10 min, an ascorbic acid solution (~4 mg in 1 mL H2O) was injected into the flask. Then, the mixture was further deoxygenated for 20 min. 5 mL of initiator modified protoplast (wet weight ~50 mg/mL) was preplaced in a 50 mL round bottom flask and then deoxygenated for 20 min with nitrogen. The polymerization solution was then transferred to the protoplast flask to initiate the polymerization. The polymerization was conducted at room temperature for a certain time and stopped by exposing the solution to air. The pellet was collected and stored at 4 °C after thorough


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centrifugation and washing steps. Characterization of polymer modified protoplasts The protoplasts of E.coli at every stage (protoplast of E.coli, the polymer modified protoplast before and after DTT treatment) were allowed to adhere onto 0.2% 3aminopropyltrimethoxysilane-coated glass/silicon slices overnight, then fixed with 3% glutaraldehyde in the dark for 2 h at 4 °C. The slides were then dehydrated in a graded ethanol series with different concentrations: 30, 40, 50, 60, 70, 80, 90 and 100% (five min per change). Then, the slides were immersed in equivalent ethanol/isoamyl acetate for 10 min and then in isoamyl acetate for 10 min, and this process was repeated once. The samples were dried and stored in a freeze dryer before SEM characterization. The morphology of the cells coated surface was characterized by SEM (Nova, NanoSEM) and confocal microscopy (A1, Nikon) using a 60 × objective. To cleave the polymer from the cell surface, the engineered cells were treated with 100 mM DTT for 5 min to cleave the disulfide bond between the cells and the polymer chain. After centrifugation, the supernatant was lyophilized and dissolved in DMF for GPC analysis (P230, Elite, China). Samples were eluted at a flow rate of 1.0 mL/min with DMF using a PolarGel-M column (7.5 mm I.D *30 cm, Agilent, USA) at 25 °C. Polystyrene standards were used for calibration. Viability and proliferation assay Wild-type E.coli BL21 cells and the protoplasts of E.coli at every stage were diluted to a final equal OD600 in autoclaved LB media. With an inoculation ratio of 1%, the diluted cell suspensions were incubated at 25 °C or 37 °C, respectively, while shaking at 200 rpm. The OD600 of the suspensions at predetermined time points was then surveyed by UV-visible spectroscopy to set up a cell growth curve. For the cell plating assay, Wild-type E.coli BL21 cells, and the protoplasts of E.coli at every stage were suspended in P buffer at a concentration of ~50 mg/mL, and plated on LB solid plates. The cells were incubated overnight at 25 °C or 37 °C, respectively. For the



Durham tube test, E.coli cells at different stages were incubated at 37 °C in a dextrose fermentation medium with bromothymol blue, under anaerobic conditions. The production of gas and the colour change were recorded. Cell membrane protein extraction by thermoprecipitation The polymer modified protoplasts were resuspended in 0.01 M PBS and broken thoroughly by ultrasonication for 30 min at 4 °C by an ultrasonic homogenizer (240 W, JY92-IIDN, Scientz, China). The supernatant was collected after centrifuging at 1200 ×g for 30 min at 4 °C. For the membrane protein separation procedure, the supernatant was further incubated in a water bath for 5 min at 40 °C, and the transparent sample became white and opaque. The white precipitate was immediately centrifuged at 7863 ×g for 10 min at 37 °C. The collected pellet was washed three times with PBS, then dissolved in 0.2% (W/V) DTT in 10% (W/V) trichloroacetone acetone solution for 4 h at -20 °C. After further centrifugation (12000 ×g, 30 min, 4 °C), the precipitate was again dissolved in 0.2% DTT in a glacial acetone solution, centrifuged (12000 ×g, 30 min, 4 °C), and dried under nitrogen gas. Finally, the sample was homogenized in lysis buffer (7 M urea, 2 M thiourea, 4% CHAPS, 1% DTT, and 2% pharmalyte). After centrifugation to remove insoluble residue (12000 ×g, 5 min, twice), the protein supernatant was collected and frozen at -80 °C before further analysis. Gel electrophoresis and MS analysis The samples were dissolved in loading buffer and run on a 10% polyacrylamide gel, followed by staining with Coomassie blue. The protein band of interest was picked and digested to peptides by trypsin. The peptides were analyzed on nanoLC-QE (EasynLC 1000 and Q Exactive, Thermo Fisher). The proteins were identified using the Mascot search engine (V. 2.2, Matrix Science, UK) and submitted to the SWISS2DPAGE database (https://world-2dpage.expasy.org/swiss-2dpage/) to annotate the location of the proteins. The 2-DE procedure was executed by the following procedure: an IPG strip with a linear pH gradient from 3 to 10 was first rehydrated in swelling


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buffer for IEF, then treated with strip equilibration buffer for 30 min. The equilibrated strips were then set up for separation in a 12% polyacrylamide gel. Proteins on the gel were visualized using Coomassie blue staining. Protein spots of interest in the gel interests were picked and digested into peptides with trypsin. Peptide analyses were performed on a 5800 MALDI-TOF/TOF (AB SCIEX). The peptides were identified using the Mascot search engine. Several possible chemical residues by polymer modification (C8H7NOS2, C3H4OS, and C9H13BrO3S2) were allowed as the variable modification for identification. Amino acid sequences of detected proteins were retrieved from http://www.UniProt.org/ and submitted to PDB (http://www.rcsb.org/) to determine the possible modification sites of the proteins. RESULTS AND DISCUSSION Polymer modification on the cell surface. Bacterial protoplast from genetically engineered E.coli was chosen for membrane protein enrichment and isolation. Compared with mammalian cell strains, bacteria are tougher and stronger and therefore more likely to retain functionality after the chemical modification process. The protoplast was formed by lysozyme treatment of the E.coli cells to expose the plasma membrane proteins. As illustrated in Scheme 1, the protoplast was reacted with a disulfide containing linker (SPDP) to selectively introduce the free sulfhydryl group to the amino groups of the membrane proteins. The ATRP initiator was then attached to the protein through a thiol-disulfide exchange reaction. A biocompatible activator regenerated by electron-transfer atom transfer radical polymerization (ARGET ATRP)30, 31 was carried out from the cell surface. Poly(Nisopropylacrylamide) (pNIPPAm), a thermoresponsive polymer, was adopted to grow on the surface of membrane proteins. The pNIPPAm tag could reversibly precipitate above the lower critical solution temperature and separate the polymer linked membrane proteins from the cytosol and subcellular organelles. The pNIPPAm tag could be further released by the cleavage of the disulfide linker with DL-dithiothreitol (DTT).



Lysozyme was selected as a model protein for verifying the efficiency and specificity of the separation strategy. Lysozyme pre-modified with initiators was mixed in crude E.coli lysate and subjected to the polymerization, thermoprecipitation, and separation steps. As shown in Figure 1, the precipitation band (Lane 2) showed a smear at the higher molecular weight, which suggested a broad polymer chain distribution tethered with lysozyme. The highly abundant protein from E.coli was almost removed from the lysozyme. After the DTT reduction, the lysozyme concentrated to be a single band at 14 kD. The light band at ~27 kD indicated that part of the protein might dimerize via a disulfide linkage. The results demonstrated that a single protein could be successfully modified with pNIPPAm, separated from the other proteins and finally purified to electrophoretic pure after the cleavage of pNIPPAm. We further investigated the possible modification site on the lysozyme by MALDI-TOF analysis. Since the SPDP reaction with the amino or guanidyl group was random, eight amino acids of lysozyme were identified as the possible sites for polymer modification (Table S1). Previous work on protein modification showed that the initiator anchoring and polymer growing procedures have little effect on the protein activity (Papain and Bromelain).29 For the cell modification, to avoid potential cell injury, the modification steps were performed in pH 7.4 PBS buffer for a limited reaction time. Field emission scanning micrographs of the protoplasts at every modification stage were taken. The results revealed that the morphologies within a strain are variable from near-spherical to a cylindrical as an individual or a chain of fused spindle-shaped cells. No distinct morphological differences were observed among the cells at the different stage of polymer modification, indicating that there was little disturbance of the cell morphology during chemical modification, polymerization, and polymer cleavage (Figure 2a-c, Figure S1). Moreover, there were no obvious cell fragments on the surface, indicating that most cell membranes still maintained their integrity. To evaluate and verify the cell viability after polymer modification, the protoplasts were recultured in growth media. Carbohydrate fermentation of the modified protoplasts was evaluated by Durham tube test (Figure 2d). All of the four samples showed a similar


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capacity to produce gas and acid from saccharide fermentation, indicating that the chemical process had little influence on the metabolic status of protoplasts. Furthermore, typical growth curves for cells at every modification stage were obtained (Figure 2e and f). The lag growth stage of the protoplast extended from 0-4 hour to 010 hour after polymer modification at 37 °C. The exponential growth patterns for the polymer modified groups were parallel with the control protoplasts. All of the groups reached a stationary stage with a similar maximum biomass accumulation. These results indicated that the protoplasts after modification were still viable, while the proliferation capability was partly inhibited by the chemical modification. The same deduction could also be drawn from the spread plate culture assay (Figure S2). It was also interesting to compare how the cell growth was influenced by the thermoresponsible pNIPPAm brushes. At 25 °C below the LCST, the polymer modified protoplasts reached the exponential stage earlier than the polymer cleaved group (30 h vs 34 h). However, when at 37 °C above the LCST, the result was the opposite. The polymer cleaved protoplasts showed a shorter lag growth stage, compared with the polymer modified group (8 h vs 10 h). We concluded that the polymer brushes became insoluble above the LCST, which may then collapse and shield the cell surface. Since the polymers grown from the membrane proteins do not cover the entire cell surface, the permeability of the cell membrane was partly affected. These results also validated that the pNIPPAm brushed were attached to the surface of the protoplast. The polymer growth on the cell surface was further investigated (Figure 3). After the cell surface-initiated polymerization, the pNIPPAm polymer brushes were attached to the cell surface through the disulfide linker. The cell suspension was lucent but a little cloudy below the LCST of the pNIPPAm, then became white and opaque when heating above the LCST. This comparison verified the successful growth of polymer from the cell surface. Further treatment with DTT specifically reduced the linker and released the grafted polymers on the cell surface. The resulting supernatant was collected by centrifugation and became cloudy and opaque, similar to the polymer modified cells above the LCST of pNIPPAm. Gel permeation chromatography analysis of the



polymers cleaved from the cell surface revealed a molecular weight of 273, 000 (Mw/Mn = 1.53). The board dispersity of the polymers could be due to the interference of radical species from the cells and the heterogeneous distribution of initiators on different site of membrane proteins. These results verified that the polymers were grown on the cell surface rather than in the cytoplasm. This reversible polymer modification enables flexible polymer anchoring on the cell surface for variable cell engineering. Preparation of the membrane proteins In traditional methods, the enrichment of membrane proteins always involves ultracentrifugation, a high concentration of detergent, and extensive washing steps with high salt and pH buffers. In this work, the separation was easily achieved by thermoprecipitation. After cell disruption, the polymer modified membrane fraction was precipitated and separated from the intracellular components at 37 °C. The polymer tag was then cleaved and removed by a DTT treatment and subsequent centrifugation at 37 °C to remove the polymer. The separated protein extract was subjected to a 12% SDS polyacrylamide gel (Figure 4). The polymer modified protein band showed a uniform background, which was attributed to the wide molecular weight distribution of pNIPPAm. After removing the polymer, some extracted protein bands disappeared in the gel, compared to the whole cell proteins, which indicated that the cytoplasmic proteins were selectively removed after the thermoprecipitation process. The enriched proteins were digested by trypsin and analyzed by LC-MS/MS. A total of 1825 proteins or protein groups were identified, of which 1445 were annotated (Supplementary Table S2). The distribution of proteins in the cell was classified: 5 (0.35%) were proteins from the extracellular matrix, 256 (17.72%) were outer membrane proteins, 117 (8.1%) were proteins from periplasmic space, and 658 (45.54%) were plasma membrane proteins. As a whole, 1036 (71.7%) proteins were membrane proteins, and 409 (28.3%) were cytoplasmic. The proteomic analysis validated the thermoprecipitation method with acceptable reliability for membrane protein enrichment and separation. The selectivity of membrane proteins enrichment was in


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agreement with or even higher than recent membrane protein enrichment studies (Table 1). Compared with the microbead tag for enrichment, 11,13 the nanoscale “grafting from” polymer tag may be less affected by the steric hindrance on the restricted membrane. Therefore, we anticipate that the nanoscale enriching strategy may have the potential to identify more complete membrane proteins. A significant proportion of the identified proteins were cytoplasmic proteins. Some of the protein original located in cytoplasm might transfer and combine with the membrane proteins. These proteins deserve further research to confirm their functionality at the plasma membrane. However, a number of the contamination proteins were induced by the strong noncovalent interactions with membrane proteins or the polymer networks. We anticipate that thorough dialysis and harsh washing could improve the selectivity, but might cause a loss of lower abundance membrane proteins. The enrichment method could be further combined with other strategies like density centrifugation to improve both the selectivity and sensitivity of membrane protein separation. To identify individual proteins separated by the thermoprecipitation method, we further subjected the extracted proteins to two-dimensional electrophoresis and compared to the whole proteins from the original strain (Figure 5). Most of the protein spots disappeared after the polymer modification and subsequent separation, which was in accordance with the SDS results. In addition, a few new spots on the separated protein gel were observed. We speculated that most of the spots on the extracted protein gel came from the outer membrane of E. Coli. However, due to experimental limits, we believe that a number of specific membrane proteins, particularly proteins with high hydrophobicity and molecular mass, were precipitated and lost during the 2-D electrophoresis process. 19 protein spots from the extracted protein gel were selected and subjected to digestion with trypsin. The resulting peptides were submitted for mass spectrometry analysis. The MS/MS spectra were identified by the Mascot search engine. Possible chemical



residues were allowed as variables for the mass fingerprint search and computation. 12 possible proteins were identified as a result (more than three peptides matched the data, and the protein score was > 95). Among them, spots 1-6 were new emerging proteins, while spots 7-12 were proteins consistent with the original strain control gel. The identified proteins were classified according to the UniProt database. Among the 12 identified Proteins, eight were annotated as membrane proteins and four were cytosolic (Table 2). The results suggested that 66.7% of the proteins separated were membrane proteins, which was consistently well matched with the proteomic results (Figure 4b), indicating good selectivity by our developed method for membrane protein analysis. Due to the limited information integrity of the E-coli strain, only three protein structures could be retrieved from the protein database: Maltoporin, Nucleosidespecific channel-forming protein TSX, and Outer membrane protein N. Most of the possible modification sites on the three proteins were located in the internal channels of the proteins (Figure S4, Table S3), which are highly associated with molecule transport. The results indicated that the chemical modification was likely to react to the lysine and arginine group in the hydrophilic barrel with little steric hindrance. Conclusion In summary, we developed a strategy involving the cytocompatible modification of thermoresponsive polymers on living cell surfaces for membrane proteomic isolation and analysis. The grafted polymer chains could be directly initiated and grown from the cell surface without losing cell viability. This strategy was highly selective as well as convenient, compared with traditional methods. The chemically selective thermal precipitation strategy could be a powerful technique for proteomic isolation and analysis of membrane proteins. AUTHOR INFORMATION Corresponding authors


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* E-mail: [email protected], [email protected] * E-mail: [email protected] Author contributions †

These two authors contributed equally to this work.

ORCID Yuanzi Wu: 0000-0003-3523-8505 Notes The authors declare no competing financial interest. ACKNOWLEDGMENTS We acknowledge the National Natural Science Foundation of China (21504105), Funds of Joint Plan for Health-Education in Fujian (83017000), Start-up Funds of Minjiang Scholar (510246), for the financial support.

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(22) Geng, W.; Wang, L.; Jiang, N.; Cao, J.; Xiao, Y. X.; Wei, H.; Yetisen, A. K.; Yang, X. Y.; Su, B. L. Nanoscale 2018, 10, 3112-3129 (23) Teramura, Y.; Kaneda, Y.; Totani, T.; Iwata, H. Biomaterials 2008, 29, 1345-1355. (24) Teramura, Y.; Iwata, H. Soft Matter. 2010, 6, 1081-1091. (25) Rossi, N. A. A.; Constantinescu, I.; Brooks, D. E.; Scott, M. D.; Kizhakkedathu, J. N. J. Am. Chem. Soc. 2010, 132, 3423-3430. (26) Zhang, P. P.; Bookstaver, M. L.; Jewell, C. M. Polymer 2017, 9, 40. (27) Cobo, I.; Li, M.; Sumerlin, B. S.; Perrier, S. Nat. Mater. 2015, 14, 143-159. (28) Wang, G.; Zhang, K.; Wang, Y. D.; Zhao, C. W.; He, B.; Ma, Y. H.; Yang, W. T. Chem. Commun. 2018, 54, 4677. (29) Wu, Y. Z.; Cai, Z.; Wu, S. G.; Xiong, W. L.; Ma, S. Y. Biopolymers 2018, 109, 23222. (30) Min, K.; Gao, H. F.; Matyjaszewski, K. Macromolecules 2007, 40, 1789-1791. (31) Matyjaszewski, K.; Jakubowski, W.; Min, K.; Tang, W.; Huang, J. Y.; Braunecker, W. A.; Tsarevsky, N. V. P. Natl. Acad. Sci. USA 2006, 103, 15309-15314. (32) Schindler, J; Lewandrowski, U.; Sickmann, A.; Friauf, E. J. Proteome Res. 2008, 7, 432-442. (33) Pischedda, F.; Szczurkowska, J.; Cirnaru, M. D.; Giesert, F.; Vezzoli, E.; Ueffing, M.; Sala, C; Francolini, M; Hauck, S, M; Cancedda, L; Piccoli, G. Mol. Cell. Proteomics. 2013, 13, 733–748.



Scheme 1. Schematic diagram of the biocompatible polymer modification of the cell membrane proteins, and the thermoprecipitation strategy for membrane protein enrichment.

Figure 1. Verification of Chemo-selective separation of lysozyme from E.coli lysate: the SDS-PAGE analysis of the mixture of lysozyme-initiator and E.coli lysate (lane 1); lysozyme-polymer conjugates after thermoprecipitation and separation (Lane 2), and recycled lysozyme after polymer cleavage (lane 3).


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Figure 2. Characterization of cell morphology and viability of the modified protoplast cells from E.coli. Field emission scanning electron microscope images of the protoplasts (a) before and (b) after polymer modification, and (c) polymer modified protoplast treated with DTT to cleave the polymer tag. Insets are of a single protoplast cell with a magnification of 30000 ×. (d) Metabolic status of protoplast cells in saccharide fermentation medium. 1) wild-type cell, 2) pristine, 3) polymer modified, and 4) polymer cleaved protoplast, respectively. (e-f) The E.coli cell growth curve of a wild-type cell (black), protoplast (blue), polymer modified protoplast (green), and polymer cleaved protoplast (red) incubated at (e) 25 °C and (f) 37 °C, respectively. OD600 indicates the cell density, Scale bars represent mean ± SD of three samples per group.



Figure 3. Polymer cleavage confirmed polymer growth on the cell surface. a) Suspension of 1) polymer-modified protoplast, 2) pristine protoplast, 3) supernatant collected from sample 1 after DTT treatment and centrifugation, and the supernatant collected from sample 1 without DTT treatment. The samples were incubated at 25 °C and 37 °C, respectively. b) GPC analysis of the supernatant by DTT treatment and centrifugation from protoplast cells (blue) and the polymers cleaved after DTT treatment of polymer-modified protoplast cells (red).

Figure 4. Chemo-selective separation of membrane proteins from E-coli protoplasts. (a) SDS-PAGE analysis of wild-type E.coli cells (lane 1); polymer-protoplast conjugates (lane 2); isolated membrane proteins by the thermoprecipitation and separation procedure (lane 3). (b) compartment assignment of the identified proteins: the proteins in lane 3 were digested to peptides and identified by mass. Protein identification can be


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found in supplemental table S2.

Figure 5. Two-dimensional gel electrophoresis of total cellular proteins of E. coli. a) Wide-type E.coli cells; b) Isolated membrane proteins by the thermoprecipitation and separation procedure. Spots that have been identified by mass are labeled to correspond with the proteins in Table 2.



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Table 1. The selectivity of membrane proteins reported by different methods in the literature Cell

Method

Protein

Membrane

identified

protein

898

Selectivity

Ref.

526

67.3%

13

338

-

40.8%

16

1698

417

62.2%

17

586

191

36.5%

32

439

-

40%

33

2158

772

36%

18

385 PMPs

21.4%

Biotinylation and affinity NCI-H1299

enrichment by streptavidin beads

Bovine semen

Biotinylation and affinity enrichment by neutravidin agarose resin

Mouse

Biotinylation and affinity

coronal

enrichment by

brain tissue

streptavidin agarose resin

Rat brain

PEG and dextran two-

tissue

phase separation Biotinylation and affinity

Cortical

enrichment by

neuron

streptavidin agarose beads

HeLa

HeLa

Glycan moieties directed enrichment

Polydopamine coating and magnetic capture

1796

11

1316 lipidraft associate

73.3%

proteins Surface-initiated E. coli

polymerization and

1445

1036

thermoprecipitation

Table 2. Proteins identified from the gel slice of 2-DE


71.7%

This work

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No.

Name

MW

PI

Peptide

(Da)

Accession no.

no.

Protein Score %

Membrane proteins 1

Outer membrane protein A

37208

5.99

28

tr|E3PIY1|E3PIY1_EC

100

OH1 2

Nucleoside-specific channel-

33568

5.07

8

forming protein tsx 3


sp|P0A927|TSX_ECO

100

LX 38109

5.61

17

tr|W1F721|W1F721_E

100

COLX 7

Outer membrane protein N

39309

4.76

6

tr|F4NE19|F4NE19_E

100

COLX 8


40182

6.24

16

tr|F4SMF4|F4SMF4_E

100

COLX 9

Outer membrane protein X

18295

5.73

3

tr|W1FVW9|W1FVW9

100

_ECOLX 10


37120

6.23

14

tr|A0A070T3V8|A0A0

100

70T3V8_ECOLX 12

Maltoporin

49951

4.86

17

sp|A8A7D6|LAMB_E

100

COHS Other 4

50S ribosomal protein L1

24714

proteins 9.64

18

sp|B7M731|RL1_ECO

100

8A 5

50S ribosomal protein L1

24728

9.64

21

6

Uncharacterized protein

189008

5.40

49

tr|A0A062Y370|A0A0

100

62Y370_ECOLX tr|A0A1E5MDA4|A0A

98.95

1E5MDA4_ECOLX 11

Superoxide dismutase

21135

5.76

3

tr|A0A1D3U2N7|A0A 1D3U2N7_ECOLX


100



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Cytocompatible Modification of Thermoresponsive ... - ACS Publications

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