Article pubs.acs.org/ac
In Cell Footprinting Coupled with Mass Spectrometry for the Structural Analysis of Proteins in Live Cells Jessica A. Espino,† Vishaal S. Mali,‡ and Lisa M. Jones*,† †
Department of Chemistry and Chemical Biology, Indiana University-Purdue University Indianapolis, Indianapolis, Indiana 46204, United States ‡ Avon High School, Avon, Indiana 46123, United States S Supporting Information *
ABSTRACT: Protein footprinting coupled with mass spectrometry has become a widely used tool for the study of protein−protein and protein−ligand interactions and protein conformational change. These methods provide residue-level analysis on protein interaction sites and have been successful in studying proteins in vitro. The extension of these methods for in cell footprinting would open an avenue to study proteins that are not amenable for in vitro studies and would probe proteins in their native environment. Here we describe the application of an oxidative-based footprinting approach inside cells in which hydroxyl radicals are used to oxidatively modify proteins. Mass spectrometry is used to detect modification sites and to calculate modification levels. The method is probing biologically relevant proteins in live cells, and proteins in various cellular compartments can be oxdiatively modified. Several different amino acid residues are modified making the method a general labeling strategy for the study of a variety of proteins. Further, comparison of the extent of oxidative modification with solvent accessible surface area reveals the method successfully probes solvent accessibility. This marks the first time protein footprinting has been performed in live cells.
P
analysis of protein−ligand and protein−protein interactions and protein conformational change.10−13 The power of the method lies in comparison of a protein in multiple states (i.e., ligand-bound and ligand-free). Differences in modification between different states provides information on protein interaction sites and protein conformations. To date, protein footprinting methods have been used in vitro on relatively pure systems. The application of these methods for in cell analysis would provide a new platform for analyzing proteins in their native cellular environment. Further, in cell footprinting would not require any genetic manipulation of the protein, such as fusing to a fluorescent indicator protein for imaging as done in FRET. Here, we describe the extension of an oxidative-based footprinting method for in cell labeling. Oxidative-based footprinting utilizes hydroxyl radicals to modify proteins. It was first coupled with mass spectrometry by Chance and co-workers14 who utilize a synchrotron to expose proteins to high energy radiations. Oxidative-based footprinting is a good choice for in cell footprinting for two reasons. First, it is a general label that can modify several amino acids.15−17 This increases its utility for various protein systems. Second, it is an irreversible label which is important for postlabeling extraction. Prior to MS analysis, cell lysis, removal of the membrane, and proteolytic
rotein interactions are essential for the regulation of many biological processes including signal transduction and cell growth.1,2 The identification of protein interactions sites provides insight into how protein activity is modulated.3 These interactions have been successfully studied in vitro by various methods such as X-ray crystallography, NMR, and fluorescence among others. A disadvantage of in vitro methods is that they monitor the protein in isolation and do not take into account other variables present in the cellular environment that could influence interactions. In vitro studies also rely on expression and purification of proteins using recombinant DNA technology. Many proteins, in particular membrane proteins, are not amenable to purification and cannot be readily studied by in vitro methods. Studying proteins in cells takes into account the effects of molecular crowding and competing cellular interactions and allows access to a number of proteins that are difficult to purify. Biochemical in vivo methods can identify interaction networks but provide limited information on interaction sites. Biophysical methods such as in vivo fluorescence resonance energy transfer (FRET) and in vivo cross-linking coupled with mass spectrometry provide information on interaction sites owing to the distance dependence of these methods.4,5 In recent years, protein footprinting coupled with mass spectrometry has been a valuable tool for the study of protein structure. These methods use either a general6,7 or amino acid specific label8,9 to modify proteins. The coupling of these methods with mass spectrometry has led to amino acid-level © XXXX American Chemical Society
Received: May 20, 2015 Accepted: July 6, 2015
A
DOI: 10.1021/acs.analchem.5b01888 Anal. Chem. XXXX, XXX, XXX−XXX
Article
Analytical Chemistry
excimer laser had a pulse frequency of 20 Hz, laser energy of 121 mJ, and pulse width of 2.55 mm. Cells were collected in a tube containing 500 μL of 40 mM of DMTU and 40 mM of PBN. DMSO (1%) was added to the quench buffer to inhibit methionine sulfoxide reductase. Cells were labeled in technical triplicate. Control samples that had H2O2 added but were not exposed to laser photolysis were also done in triplicate. Cells were prepared for MS analysis by using a mass spectrometry sample prep kit (Thermo Scientific, Pittsburgh, PA). Briefly, cells were lysed through incubation at 95 °C for 5 min in the presence of lysis buffer. Nuclease was added to digest DNA and RNA. After centrifugation, the cell lysate was reduced with DTT and alkylated with IAA. An acetone precipitation was performed on the lysate overnight. After centrifugation, the cell lysate was digested with LysC at 37 °C for 2 h and then with trypsin at 37 °C overnight. Digestion was quenched by adding 5% formic acid. A speed vac was used to dry samples which were then resuspended in 2% acetonitrile with 0.1% formic acid. Fluorescence Imaging. Vero cells grown to confluence in a T75 flask in the presence of streptomycin/penicillin were incubated with 5 μM CellROX Deep Red reagent for 30 min. Cells were trypsinized, centrifuged, washed twice with sterile PBS, and resuspended in sterile PBS. Cells were subjected to FPOP labeling as described above. After oxidative modification, cells were fixed with 3.7% formaldehyde. Cells were places in a six channel μ-Slide VI 0.4 slide (ibidi, Verona, WI) and were imaged using an Olympus Fluoview FV1000 MPE multiphoton microscope (Olympus Corporation, Center Valley, PA) with an excitation/emission wavelength of 640/665 nm. MS Analysis. Digested samples were loaded onto a 100 μm × 2 cm Acclaim PepMap100 C18 nano trap column (5 μm, 100 Å) (Thermo Scientific, Pittsburgh, PA) with an Ultimate 3000 liquid chromatograph (Thermo Scientific, Pittsburgh, PA) at 5 μL/min. The peptides were separated on a silica capillary column that was custom-packed with C18 reverse phase material (Magic, 0.075 mm × 150 mm, 5 μu, 120 Å, Michrom Bioresources, Inc., Auburn, CA). The gradient was pumped at 300 nL/min from 10 to 45% solvent B (acetonitrile, 0.1% formic acid) for 90 min, then to 90% solvent B for 5 min, and re-equilibrated to solvent A (water, 0.1% formic acid) for 12 min. The mass spectrometry was performed on a Q-Exactive Orbitrap (Thermo-Fisher, Pittsburgh, PA). The mass spectrometer was operated in data-dependent acquisition mode controlled by the Xcalibur 2.2 software. Peptide mass spectra were acquired from an m/z range of 350−2000 at high mass resolving power. The top 25 most abundant multiply charged ions were subjected to higher-energy collisional dissociation (HCD). A charge state rejection of +1 ions and monoisotopic ion selection were employed. A dynamic exclusion of 10.0 s was used. Data Analysis. For peptide identification, all MS/MS samples were analyzed by Proteome Discoverer (Thermo Fisher Scientific, San Jose, CA) using Sequest version 1.4.1.14 (Thermo Fisher Scientific, San Jose, CA) and Mascot 2.2.06 (Matrix Science, London, U.K.). The files were searched against the SwissProt human database which contains 20,165 proteins. Sequest was searched with a fragment ion tolerance of 0.020 Da and a parent ion tolerance of 10 ppm. All known hydroxyl radical side-chain reaction products16,19 were searched as variable modification. Carbamidomethylation of cysteine was specified as a fixed modification. The FPOP modifications were distributed over different search levels to reduce the computa-
digestion need to be performed. Affinity approaches such as immunoprecipitation may also be used to obtain a protein of interest. Retention of the modification would be difficult for a reversible label after these multiple steps. Although there are multiple means to generate hydroxyl radicals,18,19 we chose to use fast photochemical oxidation of proteins (FPOP). This method, first developed by Hambly and Gross,20 employs a pulsed laser for photolysis of hydrogen peroxide (H2O2) to generate hydroxyl radicals. FPOP has been used successfully to identify protein interactions sites and protein conformational changes in vitro.21−23 Hydrogen peroxide readily crosses cell membranes via free diffusion as well as through channel proteins such as aquaporin, consequently FPOP should have the capability to oxidatively modify proteins inside live cells.24 An advantage of using FPOP for in cell labeling is for oxygen mediation. Many of the oxidation reactions require oxygen19 which may not be readily available within the cell. The addition of H2O2 elicits a response from the endogenous enzyme catalase that catalyzes the decomposition of hydrogen peroxide to water and oxygen. This reaction generates an oxygen source for the subsequent oxidation reactions. In this paper, we describe the application of an in cell FPOP (IC-FPOP) method for the successful oxidative modification of endogenous proteins in live African green monkey kidney cells (Vero cells).
■
EXPERIMENTAL SECTION Reagents and Chemicals. Dimethylthiolurea (DMTU), 30% hydrogen peroxide, dimethyl sulfoxide (DMSO), and Ntert-butyl-α-phenylnitrone (PBN) were purchased from SigmaAldrich (St. Louis, MO). Vero cells were purchased from ATCC (Manassas, VA). Dulbecco’s Modified Eagle Medium (DMEM), sterile PBS, trypsin-EDTA, CellROX Deep Red reagent, and penicillin-streptomycin were purchased from Life Technologies (Grand Island, NY). HPLC-grade solvents, acetone, fetal bovine serum (FBS), and the Pierce Mass Spec Sample Prep Kit for Cultured Cells which included iodoacetamide (IAA), dithiothreitol (DTT), trypsin, and LysC were purchased from Fisher Scientific (Thermo Fisher Scientific, Waltham, MA). Cell Viability Assay. Vero cells in DMEM media supplemented with 10% FBS were cultured in a T75 flask to 70% confluence in the presence of streptomycin and penicillin. Cells were trypsinized, centrifuged, and resuspended in sterile PBS. Cells, 450, 180, and 45 μL, were added to a 24 well plate. Either from a 200 or 100 mM stock 50, 20, and 5 μL of hydrogen peroxide was added to the 450, 180, and 45 μL of cells, respectively. Cells were incubated with hydrogen peroxide for 10 min, 4 min 30 s, or 1 min 30 s for the 500, 200, and 50 μL samples, respectively, and then quenched with 20 mM of DMTU. The CellTiter-Glo 2.0 luminescent reagent (Promega, Madison, WI) was added to each well in 1:1 volume for 10 min. The luminescence was measured using an EnVision 2102 Multilabel plate reader (PerkinElmer, Waltham, MA). An ATP standard curve from 1 nM to 1 μM was also measured on the same plate. Triplicate samples were measured for each H2O2 concentration. In Cell FPOP. Vero cells grown to 70% confluence in a T75 flask in the presence of streptomycin/penicillin were trypsinized, centrifuged, and resuspended in sterile PBS. Immediately prior to FPOP, 50 μL of 200 mM hydrogen peroxide was added to 450 μL of cells. Samples were passed through a flow tube (150 μm inner diameter silica tubing) at 54 μL/min. The B
DOI: 10.1021/acs.analchem.5b01888 Anal. Chem. XXXX, XXX, XXX−XXX
Article
Analytical Chemistry tional load. The enzyme specificity was set to trypsin with 2 missed cleavages. A target decoy database search was performed. Peptide identifications were accepted if they could be established at greater than 95.0% probability to achieve a false discovery rate (FDR) less than 1.0%. The FDR was calculated by performing two separate searches, once against the SwissProt database and against a decoy database were all of the sequences from the SwissProt database are reversed. Matches from the decoy database and the nondecoy database were used to calculate the FDR. The area of the chromatograph peaks were calculated using the Precursor Ions Quantifier node in Proteome Discoverer. For quantitation of FPOP oxidation of actin with an established method,14 the raw data were aligned and converted into a centroided peaklist file by Progenesis LC-MS (Nonlinear Dynamics, Durham, NC).25 The files were searched using MASCOT 2.2.06 against the SwissProt human database. All known hydroxyl radical side-chain reaction products16,19 were searched as variable modifications. The enzyme specificity was set to trypsin with 2 missed cleavages. The mass tolerance for precursor and fragment ions was 10 ppm and 0.020 Da, respectively. Oxidative modification was quantitated on the peptide and residue level as previously described.26 Oxidation of control samples were subtracted from laser samples to eliminate background oxidation.
Figure 1. Determination of ATP content in Vero cells following exposure to FPOP relevant peroxide concentrations. ATP levels were determined as a measure of Vero cell viability. Volumes of 500, 200, and 50 μL of Vero cells were treated with no (black), 10 mM (gray), or 20 mM (white) H2O2 for 10, 4.5, and 1.5 min, respectively followed by ATP content measurement using a CellTiter-Glo 2.0 luminescent reagent and a luminometer. Values are shown as averages plus or minus the standard deviation (n = 3). No statistically significant differences were found between 10 and 20 mM peroxide as determined by student’s t test.
RESULTS AND DISCUSSION Viability of Cells in the Presence of Hydrogen Peroxide. FPOP requires the addition of hydrogen peroxide to protein samples for the generation of hydroxyl radicals via photolysis. Hydrogen peroxide is known to be toxic to cells in a concentration- and time-dependent manner.27 Significant cell death upon the addition of H2O2 would reduce the efficacy of IC- FPOP. To determine the viability of cells after exposure to H2O2 in the time frame of an IC-FPOP experiment, the ATP content of peroxide treated Vero cells was measured. For in cell FPOP, cells are under constant flow (54 μL/min) toward the laser. The time it takes for an entire sample to reach the laser is dependent on sample volume. Three volumes of cells, 500, 200, and 50 μL, were treated with peroxide for 10, 4.5, and 1.5 min, respectively, to demonstrate the effect of time of exposure. Cell suspensions of 500, 200, and 50 μL contain 1.05 × 106, 4.2 × 105, and 1.05 × 105 cells, respectively. Two different peroxide concentrations, 10 and 20 mM, were also tested. After incubation, H2O2 was quenched with dimethyl-thiourea (DMTU), a cell-permeable H2O2 quencher, and the concentration of ATP was measured using luminescence. The ATP content correlates with luminescence intensity, the stronger the luminescence signal the higher the concentration of ATP. Upon the addition of 10 and 20 mM hydrogen peroxide, the ATP content (measured as luminescence intensity in counts/ second) decreases at all times indicating some cell death (Figure 1). However, at all cell volumes, the decrease in ATP is
