Immunopharmacology and Quantitative Analysis of Tyrosine Kinase Signaling

Abstract In this article we describe the use of pharmacological and genetic tools coupled with immunoblotting (Western blotting) and targeted mass spectrometry to quantify immune signaling and cell activation mediated by tyrosine kinases. Transfer of the ATP γ phosphate to a protein tyrosine residue activates signaling cascades regulating the differentiation, survival, and effector functions of all cells, with unique roles in immune antigen receptor, polarization, and other signaling pathways. Defining the substrates and scaffolding interactions of tyrosine kinases is critical for revealing and therapeutically manipulating mechanisms of immune regulation. Quantitative analysis of the amplitude and kinetics of these effects is becoming ever more accessible experimentally and increasingly important for predicting complex downstream effects of therapeutics and for building computational models. Secondarily, quantitative analysis is increasingly expected by reviewers and journal editors, and statistical analysis of biological replicates can bolster claims of experimental rigor and reproducibility. Here we outline methods for perturbing tyrosine kinase activity in cells and quantifying protein phosphorylation in lysates and immunoprecipitates. The immunoblotting techniques are a guide to probing the dynamics of protein abundance, protein–protein interactions, and changes in post‐translational modification. Immunoprecipitated protein complexes can also be subjected to targeted mass spectrometry to probe novel sites of modification and multiply modified or understudied proteins that cannot be resolved by immunoblotting. Together, these protocols form a framework for identifying the unique contributions of tyrosine kinases to cell activation and elucidating the mechanisms governing immune cell regulation in health and disease. © 2020 The Authors. Basic Protocol 1: Quantifying protein phosphorylation via immunoblotting and near‐infrared imaging Alternate Protocol: Visualizing immunoblots using chemiluminescence Basic Protocol 2: Enriching target proteins and isolation of protein complexes by immunoprecipitation Support Protocol: Covalent conjugation of antibodies to functionalized beads Basic Protocol 3: Quantifying proteins and post‐translational modifications by targeted mass spectrometry


INTRODUCTION
Tyrosine kinases are critical mediators of immune cell activation and regulation (Hwang, Byeon, Kim, & Park, 2020;Lowell, 2011). The transfer of the ATP γ phosphate to a protein tyrosine residue initiates signaling cascades that alter cell survival, proliferation, and effector functions. The steric and electrostatic effects of tyrosine phosphorylation can induce conformational changes in proteins that expose docking sites, block autoinhibitory interactions, or deprotect motifs for trafficking, degradation, or further post-translational modification. Phosphotyrosine-containing peptides also serve as direct SH2 and PTB binding sites, nucleating higher-order signaling complexes that tune signal strength and kinetics and may even alter the phase properties of signaling complexes (Case, Ditlev, & Rosen, 2019;Oh et al., 2012). The actions of tyrosine kinases initiate an array of immune cell functions, including pathogen detection and killing, phagocytosis, clonal expansion, and migration to sites of infection or damage.
Accordingly, dysregulation of tyrosine kinase signaling pathways is associated with many diseases, including autoimmune and inflammatory disease and cancer. Analysis of activated signaling pathways, therefore, is critical for understanding how immune cells participate in health and disease.
In this article we highlight genetic and chemical tools-including competitive inhibitors, designer kinase-inhibitor pairs, small interfering RNA (siRNA), and CRISPR/Cas9 gene editing-for dissecting tyrosine kinase signaling in immune cells. We present protocols for quantitative evaluation of signaling kinetics, amplitude, and binding interactions and for identifying sites of post-translational modification. Our protocols feature adherent bone marrow-derived macrophages (BMDMs), but we describe adaptations for use with lymphocytes and other cells in suspension. Basic Protocol 1 describes a method for quantitative immunoblotting. Basic Protocol 2 describes a method for (co-)immunoprecipitation of proteins from cell lysates, which can be used in conjunction with immunoblotting or quantitative, targeted mass spectrometry described in Basic Protocol 3. These cell perturbation and protein enrichment strategies can also be used as precursors to flow cytometry or proteomic methods (see Current Protocols articles: Breitkopf & Asara, 2012;Schulz, Danna, Krutzik, & Nolan, 2012). recognition of epitopes and subsequent coupling to a luminescent readout. This protocol contains instructions for quantification of total protein and phosphoprotein content with near-infrared imaging of fluorophore-conjugated secondary antibodies. Near-infrared imaging (LI-COR Odyssey or equivalent) has a broad dynamic range amenable to densitometry quantification in LI-COR Image Studio Lite or other software package (e.g., NIH ImageJ; see Internet Resources). The Alternate Protocol describes visualization of blots by chemiluminescence imaging.
We describe a method for stimulating adherent BMDMs with depleted zymosan, a βglucan preparation that binds the hemi-ITAM-containing receptor Dectin-1 (Underhill, 2003). This representative cell-activating stimulus can be coupled with pharmacological, transcriptional, or genetic disruption of tyrosine kinase function to test the contribution of these kinases to cell signaling. Alternative receptor ligation or inhibition of analogsensitive Csk (Csk AS ) by the small molecule 3-IB-PP1 can be used as an alternative to Dectin-1 clustering. In the latter approach, 3-IB-PP1 inhibits a sensitized form of Csk, the tyrosine kinase that negatively regulates the Src family tyrosine kinases (SFKs). When Csk AS is inhibited, SFKs become activated and initiate signaling through many pathways (see Background Information; Brian et al., 2019;Freedman et al., 2015;Schoenborn, Tan, Zhang, Shokat, & Weiss, 2011;Tan et al., 2014). Dectin-1 ligation is a useful positive control for myeloid cell activation via tyrosine kinase-dependent signaling (Freedman et al., 2015;Goodridge et al., 2011), but the choice of controls for a given experiment should reflect the cell and pathway of interest. Where appropriate, we include adaptations applicable to lymphocytes and other cells in suspension.
8. Gently apply 0.5 ml sonicated and washed depleted zymosan or 3-IB-PP1 with or without kinase inhibitor (or alternative stimulation/perturbation). Quickly but gently place plates in the prewarmed centrifuge, and pulse spin 30 s at 5000 × g to synchronize deposition of depleted zymosan particles onto cells. with molecular weight marker. Include positive and negative controls on each gel to facilitate quantification across blots. Load unused wells with SDS sample buffer.
The loading strategy is appropriate for immunoblots from whole-cell lysates for moderately expressed proteins using near-infrared imaging. Optimization may be needed if probing extremely abundant or rare proteins, events, or immunoprecipitates.
If blots will later be cut horizontally, straight cutting may be facilitated by loading twoto three-times diluted (to indicate left to right directionality) molecular weight marker in the right-most lane. If blots will be cut vertically, it is advisable to use marker lanes between segments (Fig. 1).
16. Fill electrophoresis module with running buffer, and apply constant voltage (150 V) until the dye front has migrated out of the gel or the desired separation has occurred (∼80 min).
The time and voltage will depend on the size of the target protein(s) and the gel and buffer system being used.
This protocol uses wet transfer methodology, which generally produces the highestquality results across a wide range of molecular weights. See manufacturer's instructions for semi-dry transfer buffers and setup.
18. Assemble transfer apparatus according to the manufacturer's instructions. Place one corner of the membrane on top of the gel. Slowly place the opposite corner of the membrane onto gel, and lower the rest of the membrane onto the gel, taking care to avoid trapping bubbles. Orient transfer with the membrane on the positive (anode) side and gel on the negative (cathode) side.
19. Fill inner and outer chambers of the apparatus with cold transfer buffer. Place on ice or in a cold room.
Chilling is critical for minimizing heat deformation of the gel during transfer.
The timing and voltage may need to be optimized.

21.
Remove membrane from the apparatus, and dry in between two sheets of clean filter paper to fix proteins onto the membrane. 24. Discard total protein stain, and wash two times for 30 s each with total protein wash.
25. Rinse membrane three times with water, and image gel with a near-infrared imaging system.
26. Rinse membrane briefly in water. Replace solution with total protein removal solution. Rock 5 min in the dark at room temperature.
Immunoblotting 27. Discard solution and place membrane in methanol.
28. If cutting membrane into segments of different molecular weights, place membrane on clean filter paper, and cut with clean scissors or razor blade. Return to methanol.
29. Discard methanol and rinse three times for 30 s each with water.
30. Discard water. Rock 2 min in 5 to 10 ml TBS at room temperature.
31. Discard TBS and add 5 to 10 ml blocking buffer. Rock 1 hr at room temperature in the dark.
Blocking buffer may be purchased from commercial vendors (e.g., LI-COR)  32. Dilute primary antibody in 4 to 6 ml (depending on the size of the container) of 1:1 blocking buffer:primary diluent.
Optimal dilution will vary by antibody. Antibodies of different species can be combined in the same solution if multiple emission wavelength channels are available (e.g., rabbitderived anti-phospho-Erk1/2 combined with mouse-derived anti-Erk1/2 imaged in separate channels).
If an antibody on an uncut gel is sufficiently specific as to produce a single band in a particular experimental condition, multiple antibodies of the same color and/or species may be pooled when detecting proteins separated by molecular weight. If there is any doubt about specificity, cut blots instead of combining antibodies.
An optimal dilution of primary antibody should be evaluated by titration for each cell type and stimulation condition, but a good starting range is 1:1000 to 1:5000. With nearinfrared imaging, primary antibodies can typically be diluted 2 to 20 times more than recommended by the manufacturer.

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Current Protocols in Immunology The software will take box size into account, but it is best to keep the boxes uniform. 43. In the Background pane, select User Defined for background quantification.
44. Draw a small box in between two lanes with representative background fluorescence. In the Background tab, select Assign Shape to apply this box for background subtraction.
45. Export data from the Shapes tab into Microsoft Excel or other spreadsheet manager. Use the background-corrected "Signal" column for data normalization and graphing. 48. In the Background pane, select Median for background quantification. Adjust borders to top/bottom or right/left, and choose the background box size.

Quantification of immunoblots (repeat for each protein of interest)
The directionality and size of background boxes will depend on the shape of the band being quantified, how well separated the lanes are, whether there are nonspecific or unidentified bands above or below the band of interest, and whether the lanes are generally higher in the background than the space in between lanes.
49. Export data from the Shapes tab into Microsoft Excel or other spreadsheet manager. Use the background-corrected "Signal" column for data normalization and graphing.
Report the abundance of a protein or modification relative to the total protein stain within the same lane. It may be appropriate to report post-translational modifications relative to a total protein immunoblot for the protein of interest. These two analytical approaches will reflect an overall dose in the cell population versus a more stoichiometry-like assessment of the degree of modification within the existing protein.
It may also be useful to perform a second normalization step relative to a reference (e.g., time = 0, wild-type, or unpolarized) value. This will obscure basal differences between treatment groups but clarify differences in kinetic response to the cell treatment or perturbation.

VISUALIZING IMMUNOBLOTS USING CHEMILUMINESCENCE
Horseradish peroxidase (HRP)-conjugated antibodies in conjunction with chemiluminescence imaging is another common approach to visualizing immunoblots. In contrast to direct dye conjugation in near-infrared imaging, HRP-adsorbed blots are developed Brian et al.

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Current Protocols in Immunology by addition of an HRP substrate (a luminol/enhancer mixture) that generates a chemiluminescent signal from HRP-conjugated secondary antibodies. Although this method can in some cases be quantitative, the dynamic range is typically narrower than in near-infrared imaging, and it is easy to over-or under-produce signal in this indirect method. To achieve the best signal, gel loading, antibody dose, and substrate choice should be optimized. An advantage of this approach is that the HRP enzyme can be efficiently inactivated and the blot reprobed with a different set of antibodies.
As in Basic Protocol 1, the dilution of each antibody must be optimized. For chemiluminescence imaging, the primary antibody (Basic Protocol 1 step 32) will typically be diluted according to manufacturer's instructions. HRP-conjugated secondary antibodies are typically diluted 1:100,000.
The HRP enzyme is sensitive to azide (N 3 ), so it is important to wash blots thoroughly after incubation with primary antibody.

As described in Basic Protocol 1, with proper controls, antibodies can be combined if their output is highly specific and/or clearly identifiable by molecular weight.
Unlike near-infrared fluorescence imaging, HRP antibody-bound membranes can be treated with NaN 3 and frozen to inactivate the HRP enzyme. Blots can then be reprobed with different antibodies (Freedman et al., 2015). This is especially useful if the second set of antibodies is derived from a different species to prevent fresh HRP secondary antibody binding to the previously adsorbed primary antibody.
Buffers are marketed for stripping blotting antibodies from membranes. In our experience this can work but often not uniformly across the membrane surface, which can limit the fidelity of quantification. We recommend performing multiple blots rather than stripping if HRP activation is not sufficient to resolve different sets of proteins.
2. Prepare substrate working solution by combining equal amounts of peroxide and enhancer solutions (from SuperSignal kit). Place membrane on a piece of plastic wrap, and pipet a minimum volume of substrate working solution onto the surface of the blot. Tilt membrane to thoroughly coat, and watch for bands to develop, following manufacturer's instructions.
HRP working solution is stable for ∼8 hr and can be reused.
3. Cover membrane in clear plastic, and smooth to remove bubbles. Image using a luminescence imaging system.

ENRICHING TARGET PROTEINS AND ISOLATION OF PROTEIN COMPLEXES BY IMMUNOPRECIPITATION
First described in the 1970s (Kessler, 1975), immunoprecipitation is a common method for separating proteins from cell lysates in denaturing or nondenaturing conditions. It has been further refined for protein purification, enrichment of low-abundance species, and identification of protein complexes (co-immunoprecipitation). As a tool for studying cell signaling, immunoprecipitation typically starts with preparation of an antibody-bead complex (noncovalent in this protocol, covalent in the Support Protocol). These antibody-coated beads are then mixed with cell lysates and gently tumbled under conditions that maximize protein capture but minimize protein degradation and further post-translational modification. The protein-antibody-bead complex is then collected, and the (co-)immunoprecipitated proteins are eluted for analysis (Fig. 2).
This protocol describes immunoprecipitation of a target protein from cell lysate. In the absence of the usual loading controls available in whole-cell lysate (described in Basic Protocol 1), a protein content normalization step is essential for quantitative analysis. The immunoprecipitation time, detergent, and salt content will determine the extent of interacting protein co-immunoprecipitation. The composition of the immunoprecipitate can then be probed by blotting for pan-phosphotyrosine, specific phosphorylation sites, total protein, or other epitopes to reveal protein-protein interactions and post-translational modifications that follow cell perturbation (as in Basic Protocol 1). The immunoprecipitates can also be subjected to phosphoproteomics to identify unknown proteins or targeted mass spectrometry to quantify specific peptides and post-translational modifications. A targeted approach is described in Basic Protocol 3.

Cell stimulation and lysis
1. Collect 4 to 5 μl protein G Sepharose beads per 10 6 cells by microcentrifuging 2 min at 500 × g, room temperature. Carefully remove supernatant, and replace with PBS. Pulse vortex, collect beads, and repeat wash two times.
The choice of immunoprecipitation antibody (subclass, host species, and prior functionalization) and any functionalization of the cells or lysates will determine the optimal bead adsorption modality (protein G, protein A, streptavidin, other). As an example, see Table  1 for specificities of proteins G and A.
2. Prebind immunoprecipitation antibody to beads by incubating 1 to 2 μg antibody per 40 × 10 6 cells with beads. Rotate beads at least 2 hr at room temperature prior to use.
In this protocol the immunoprecipitation antibody will co-elute with the target protein, possibly yielding dark bands caused by the antibody heavy chain at ∼50 to 70 kDa and the light chain at 25 kDa (Harlow & Lane, 1988). Secondary antibodies may even react across species owing to the sheer abundance of the antibody bands. For optimal visualization and quantification of proteins close to either molecular weight, it is advisable to Brian et al.

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Current Protocols in Immunology use covalent conjugation to prevent antibody elution from the beads. One such method is described in the Support Protocol.
3. Add protease and phosphatase inhibitors to an aliquot of chilled lauryl maltoside lysis buffer. Protect from light and keep on ice until use.
We have had the most success using lauryl maltoside detergent for cell lysis and immunoprecipitation. Other detergents, such as NP-40 alternative, can decrease the number of loosely interacting proteins that co-immunoprecipitate with the target.
NP-40 alternative is less expensive than lauryl maltoside and can be substituted in wash steps.
4. Prepare and treat cells as described in steps 1 to 9 of Basic Protocol 1.
If stimulating a large number of cells, they can be rested on larger plates (e.g., 150 mm 2 ).
5. Quench signaling by washing cells two times with ice-cold PBS and placing plates on ice.
7. Scrape plates to lift cells, and collect in sterile 1.5-ml LoBind tubes. 17. Wash beads and column five times with 1 ml NP-40 alternative wash buffer, centrifuging 2 min at 450 × g, 4°C, and discarding flow-through between washes.
To avoid disrupting protein complexes, it may be desirable to continue using lauryl maltoside or other detergent instead of NP-40 alternative.
18. Elute immunoprecipitated protein by applying enough immunoprecipitation elution buffer to cover the beads in the spin column. Incubate 15 min at room temperature.
19. Incubate lysate and immunoprecipitate samples 5 min at ≥99°C. Centrifuge samples 1 min at 10,000 × g, 4°C. Handle and store gel samples as described in step 12 of Basic Protocol 1.
20. To assess the efficiency of immunoprecipitation and the general stoichiometry of co-immunoprecipitated protein binding, run immunoblots with whole-cell and immunodepleted lysates, as described in steps 13 to 46 of Basic Protocol 1. Assess immunoprecipitates by immunoblot, skipping the total protein stain in steps 22 to 26 of Basic Protocol 1.
Immunoprecipitates can be further probed by immunoblotting as described in Basic Protocol 1 or by targeted mass spectrometry as described in Basic Protocol 3. These samples can also be analyzed using unbiased mass spectrometry.

COVALENT CONJUGATION OF ANTIBODIES TO FUNCTIONALIZED BEADS
In some cases, it is best to conjugate immunoprecipitation antibodies covalently to immunoprecipitation beads rather than co-eluting antibodies with the immunoprecipitate samples. For immunoblotting analysis (Basic Protocol 1), covalently conjugated antibody-beads complexes produce cleaner images, facilitating visualization and quantification of proteins comigrating with the heavy and light chains. Covalent conjugation is also ideal for subsequent mass spectrometry analysis (Basic Protocol 3) in that resulting immunoprecipitates can be desalted and run directly without further purification of proteins or detergents that would otherwise harm the mass spectrometer. In spite of these advantages, covalent conjugation tends to be used selectively because of the increased investment of time and reagents.
1. Collect enough protein G Sepharose (or alternative) beads for each conjugation.
Each vial of DMP yields 10 ml crosslinking solution, enough for one conjugation reaction with 1 ml beads. The antibody and beads should be titrated for the specific immunoprecipitation. Start with 2 μl antibody and 50 μl beads per 8 × 10 6 macrophages.
As with immunoblotting, start by doubling the number of lymphocytes or other cells in suspension to ensure adequate material for immunoprecipitation.
2. Wash beads twice with PBS by centrifuging 30 s at 1000 × g, room temperature.

Resuspend in PBS.
4. Add immunoprecipitation antibody, and tumble 1 hr at room temperature. 10. Spin beads down by briefly centrifuging and remove ethanolamine. Elute unbound antibody by incubating two times for 10 min each with 1 M glycine, pH 3.0, at room temperature.

QUANTIFYING PROTEINS AND POST-TRANSLATIONAL MODIFICATIONS BY TARGETED MASS SPECTROMETRY
Commercial antibodies are not available for every protein epitope and post-translational modification. Antibodies may be raised against custom sequences, but this process is costly and at times problematic. Mass spectrometry is a valuable tool for detecting changes in protein homeostasis and identifying novel sites of modification prior to investing in antibody generation. We present a protocol for quantifying phosphorylation on an immunoprecipitated protein via targeted liquid chromatography-tandem mass spectrometry (LC-MS/MS), using parallel reaction monitoring (PRM) on a high-resolution mass spectrometer. Traditional, data-dependent acquisition triggers fragmentation of the Brian et al.

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Current Protocols in Immunology PRM instead uses precursor ion selection to trigger fragmentation of modified peptides of interest and creating full-scan MS2 spectra, improving selectivity, sensitivity, and signal-to-noise ratios (Rauniyar, 2015). This protocol may be adapted for other epitopes or modifications and for kinase (or other) activity assays for probing modification of a target site in cells, lysates, or recombinant proteins.
We describe steps to ensure accurate peptide identification and quantification using a heavy isotope-labeled internal standard. Prior to beginning proteolytic in-gel digestion, a BCA assay is used to quantify the total protein concentration in cell lysates, a critical step for normalizing phosphopeptide levels across samples. Subsequently, protein concentrations are determined using a standard curve of titrated BSA, quantified by densitometry after SDS-PAGE. This ensures that phosphopeptide quantification can be expressed as a concentration ratio relative to the amount of protein subjected to tryptic digest. Finally, a stable isotope-labeled reference peptide is spiked into the immunoprecipitate prior to proteolytic digestion. Peptide concentrations can then be definitively identified and placed on an absolute scale via a peptide standard curve (Fig. 3).
2. Quantify amount of immunoprecipitated protein for each sample via densitometry (Basic Protocol 1). Create a standard curve using signals from the serially diluted lanes to calculate the relative amount of precipitated protein in each sample.
3. Concentrate equal amounts of immunoprecipitated protein sample with as much lysate as possible to ensure detection of potentially rare peptides via LC-MS/MS and using centrifugal filter columns according to manufacturer's instructions. Store eluent at −80°C indefinitely.

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Current Protocols in Immunology

Gel electrophoresis and quantification
For the following steps, wear a face mask and gloves to minimize keratin contamination during gel loading, excision, reduction, and alkylation.
4. Prepare BSA protein quantification standards at 0.05 to 20 μg total per lane. 9. Generate a standard curve via densitometry analysis of the BSA bands, as described in Basic Protocol 1 and shown in Figure 4. Use this curve to quantify the amount of experimental sample in each lane.

We typically quantify the total protein in each lane across all molecular weights. It may be more appropriate with recombinant proteins to quantify only the band of interest in each immunoprecipitate.
10. Using a fresh, clean razor blade for each sample, excise a sample of gel corresponding to the desired protein (molecular weight range), and place into 1.5-ml LoBind tubes.

Reduction and alkylation
11. Cut gel samples into small (∼2 mm) pieces. Wash gel fragments three times for 15 min each by submerging in ∼100 μl (depending on gel fragment size) of a 1:1 mixture of 100 mM aqueous ammonium bicarbonate:acetonitrile. Mix by vortexing prior to each incubation.
12. Remove final wash, and incubate 1 min in 100% acetonitrile, until gel pieces turn opaque. Collect fragments by briefly centrifuging in a microcentrifuge and discard acetonitrile.
13. Submerge gel fragments in an aqueous solution of 10 mM DTT/50 mM ammonium bicarbonate. Incubate 1 hr at 56°C. Pulse spin in a microcentrifuge and remove supernatant.

Make DTT solution fresh by dissolving DTT into 50 mM ammonium bicarbonate.
14. Submerge fragments in an aqueous solution of 55 mM iodoacetamide/50 mM ammonium bicarbonate. Incubate 30 min at room temperature in the dark. Pulse spin in a microcentrifuge and remove supernatant. 15. Wash gel fragments twice with a 1:1 mixture of 100 mM ammonium bicarbonate:acetonitrile.
16. Remove solution and dry fragments by incubating 1 min in 100% acetonitrile.
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Current Protocols in Immunology In-gel protease digest 17. Remove acetonitrile and cover with trypsin digest solution (see Shevchenko, Wilm, Vorm, & Mann, 1996) spiked with 13 C, 15 N-heavy isotope amino acid-labeled reference peptides. Incubate 15 min on ice.
The precise concentration of heavy-labeled reference peptide spiked in during in-gel digest will depend on the final yield of protein extracted from the gel. This can be estimated by densitometry from the BSA curve generated after SDS-PAGE. However, we suggest doing a trial run to ensure the spiked-in reference peptide is not orders of magnitude higher or lower in concentration than the peptides of interest. Trypsin, which cuts at lysine and arginine residues (Ma, Tang, & Lai, 2005), is often the protease of choice. If the distribution of lysine and arginine around the sequence of interest is suboptimal for LC-MS/MS detection, another protease such as chymotrypsin, LysC, or LysN (Giansanti, Tsiatsiani, Low, & Heck, 2016)  24. Remove solvent by vacuum concentration (e.g., SpeedVac), and store at −80°C indefinitely.

Preparation of calibration curve samples
32. Dilute 1000 fmol heavy isotope-labeled phosphorylated peptide standard in calibration curve buffer into several LoBind tubes.
33. To create a calibration curve to quantify the amount phosphorylated peptide, add increasing concentrations of unlabeled phosphorylated peptide standard so that the molar ratio of unlabeled phosphorylated peptide standard:heavy-labeled phosphorylated peptide standard spans 0.1 to 1.5.
The precise molar ratios will depend on the assay and the amount of phosphorylated peptide in each sample.
34. Dilute 1000 fmol phosphorylated peptide standard in calibration curve buffer into several LoBind tubes.
35. To create a calibration curve to quantify the ratio of phosphorylated and unphosphorylated peptide in each sample, add increasing amounts of unphosphorylated peptide standard such that the molar ratio ranges from 0.05 to 1.5.
The peptide ratios used in this calibration curve will depend on the stoichiometry of tyrosine phosphorylation and may need to be adjusted depending on the rarity of phosphorylation for a given phosphorylation site.
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Current Protocols in Immunology 36. Submit calibration curve samples and in-gel digested samples for LC-MS/MS (steps 37 to 42).
38. Mount loaded PicoTip Emitter column in a nanospray source in line with an Orbitrap Fusion with 2.1 kV spray voltage in the positive mode and heated capillary maintained at 275°C.
39. Set up a tripartite peptide elution program decreasing the fraction of HPLC buffer A and increasing the fraction of HPLC buffer B with a 300 nl/min flow rate: 5% to 10% HPLC buffer B over 5 min 10% to 16% HPLC buffer B over 40 min 16% to 26% HPLC buffer B over 5 min.
The elution program should be optimized depending on the m/z and hydrophobicity of the target peptide and desired resolution. This step presents a general starting point in three gradient stages.
40. Define an acquisition method comprising a full scan and PRM to detect singly, doubly, and triply charged precursor ions without scheduling. Set the full scan event to employ a 380 to 1500 m/z selection, an Orbitrap resolution of 60,000 (at m/z 200), a target automatic gain control (AGC) value of 4 × 10 5 , and maximum ion injection time of 50 ms. Set the PRM scan to employ an Orbitrap resolution of 30,000 (at m/z 200) and a target AGC value of 5 × 10 4 and/or maximum ion injection time of 54 ms to ensure that enough fragment ions are captured for MS/MS detection.
The acquisition method and scan events will vary depending on the chemical composition of the targets and the number of peptides analyzed in a given experiment. When there are few peptides, method development can be simplified by monitoring selected precursor ions for the duration of the chromatography gradient. For quantitative experiments, a selected peptide must be surveyed and an MS2 acquired at least 10 times across the extracted ion chromatogram (EIC). Scheduling may be used when the number of possible peptide precursors is >20 in order to capture 10 MS2 scans across a peptide chromatogram.
Quantification can be performed using MS1 or MS2 (the two components of MS/MS) EICs in the Skyline software package (see Internet Resources). A full spectrum scan facilitates assessment of dynamic range issues, co-eluting peptides, and complexity of the sample and acts as an additional validation of accurate mass for the peptide of interest. The m/z range defined above surveys all ions with a charge >1. At the above resolution EICs can be used to uniquely identify co-eluting peptides with small m/z differences so they can be fragmented individually for identification by MS2. Fill time is simply the time we allow the ions to fill the chamber.
41. Set the MS2 quadrupole isolation window to 1.6 m/z. Perform fragmentation with a higher-energy collision-induced dissociation (HCD) of 30%, and collect an MS2 scan from 100 to 1000 m/z. HCD will depend on peptide chemistry and phosphorylation site sequence context, so it will have to be optimized (Diedrich, Pinto, & Yates, 2013).
42. Collect PRM data in centroid mode, and export for quantification.

Centroid data acquisition decreases file size.
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Current Protocols in Immunology Importing and inspecting standard raw data 47. Import raw mass spectrometer files into Skyline by navigating to File, Import, and Results. Choose Add single injection replicates in files and select OK, which will prompt the Import Results Files to show raw standard curve data files. Upload the selected files by choosing Open, followed by Do Not Remove when the option to remove the naming prefix appears. Confirm and close the window by selecting OK.
48. Using raw files generated from standards (e.g., heavy isotope and light isotope phosphopeptides), inspect the chromatographic traces for quality control.
If chromatographic peaks have a non-Gaussian peak shape, the samples and standards should be rerun for quality assurance. Inconsistent LC retention times could reflect inadequate chromatographic resolution. Phosphopeptide transition ions should be chosen based on relative signal intensity of their EIC and selected for ions that are representative of larger y or b ions in the peptide fragmentation series. For example, a 10-mer peptide may fragment to yield a y 9 ion, but the y 8 ion may exhibit an EIC that is higher intensity and should thus be selected for quantification. Peptide sequence and length also affect selection of transitions for peptide validation and quantification.

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Current Protocols in Immunology 54. Define the standard raw files as Sample Type Standard, and specify their Analyte Concentrations. Select Unknown for sample raw files.
55. To view the calibration curve, go to the View menu, and select Calibration Curve.
56. Access Reports from the Document Grid, and select Peptide Quantification. Prepare a report in the Export tab, enter the file name, and click OK.
57. Normalize raw quantifications for each sample using the total protein amount used for in-gel digestion.

Background Information
Tyrosine kinases are important regulators of immune cell activation, proliferation, and survival (Bryan & Rajapaksa, 2018). Transfer of the terminal phosphate of ATP to a tyrosine residue on a protein substrate results in changes in conformation and protein-protein interaction that act as signals to direct cellular Brian et al.

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Current Protocols in Immunology  (Lemmon & Schlessinger, 2010). The growth, survival, and proliferation functions of tyrosine kinases are important in all cells. Immune cells employ peculiar binding motifs, alternative expression of kinase family members, and additional receptor families for additional functionalities such as phagocytosis, antigen-specific signaling, and polarization. In lymphocytes, hematopoietic SFKs initiate signaling downstream of T and B cell receptors by phosphorylating immunoreceptor tyrosine-based activation motifs (ITAMs), which leads to activation of the tandem SH2containing tyrosine kinases Syk and Zap-70. Together, these tyrosine kinases activate FAK family tyrosine kinases (FAK, Pyk2) and Tec family tyrosine kinases (Btk, Itk, Tec; Hwang et al., 2020). Parallel pathways are activated upon Fc receptor engagement in myeloid and NK cells (Bradshaw, 2010;Cox & Greenberg, 2001;Freedman et al., 2015;Futosi & Mocsai, 2016;Lowell, 2011). Janus kinase (JAK) activation downstream of receptor tyrosine kinases is critical for activation of signal transducer and activator of transcription (STAT) proteins that mediate growth, differentiation, and polarization (Villarino, Kanno, & O'Shea, 2017). Other receptor tyrosine kinases such as Flt3, c-Kit, and Tyro/Axl/Mer control cell survival, differentiation, and many other essential functions of immune cells (Masson & Ronnstrand, 2009;Rothlin, Carrera-Silva, Bosurgi, & Ghosh, 2015). Despite the many inputs that engage tyrosine kinases and an intense research focus on the tyrosine kinases involved in immune activation, we are still discovering elements of the interactions and dynamics of tyrosine kinases with profound effects on immune regulation (Brian et al., 2019;Courtney et al., 2017;Freedman et al., 2015;Hwang et al., 2020;Salter et al., 2018). Understanding the dynamics, kinetics, substrates, and scaffolding interactions of tyrosine kinases is critical to developing therapeutics that modulate immune function (Roschewski et al., 2020;Salter et al., 2018;Solouki, August, & Huang, 2019).
Numerous tools exist for studying the actions of tyrosine kinases in immune cells, including genetic methods such as siRNA knockdown, CRISPR/Cas9-based gene editing, small-molecule inhibitors, and chemicalgenetic designer kinase-inhibitor pairs. Each approach has advantages and disadvantages with regard to specificity, temporal control, and likelihood of triggering compensatory mechanisms (Table 2).
Genetic methods are attractive options for studying kinase function because of their inherent specificity and stability. While knockout gene editing strategies are valuable because they offer complete disruption of kinase signaling, siRNAs offer inducible control over kinase signaling disruption and are especially useful when knocking out a given kinase is lethal or maturation-impairing to a cell type or animal. siRNAs and genetic knockouts are routinely used to investigate the roles of Brian et al.

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Current Protocols in Immunology kinases in immune cells. For instance, mice in which the SFK Lyn has been knocked out have become important models of autoimmune disease after studies revealed the importance of Lyn as a negative regulator of B cell and dendritic cell activation (Brodie, Infantino, Low, & Tarlinton, 2018;Scapini, Pereira, Zhang, & Lowell, 2009). Tyrosine kinase knockouts can also be coupled to Cre-lox and FLP-FRT systems for cell-specific knockout (Lamagna, Hu, DeFranco, & Lowell, 2014;Lamagna, Scapini, van Ziffle, DeFranco, & Lowell, 2013). The advent of CRISPR-Cas9 gene editing has facilitated the substitution of specific amino acid residues in knockin models, allowing researchers to dissect novel elements of tyrosine kinase function (Harder et al., 2001). The major drawback of knockout and knockdown models for studying kinase signaling is that cells often develop compensatory mechanisms for coping with loss of the given kinase. These feedback (or, in cell lines, evolutionary) effects may mask the normal signaling contributions and scaffolding interactions of the kinase of interest (El-Brolosy & Stainier, 2017;Peng, 2019).
Small-molecule inhibitors have facilitated the study of kinases in many aspects of immune activation. Kinase inhibitors generally function by competing with ATP for access to the active site, preventing substrate phosphorylation (Davies, Reddy, Caivano, & Cohen, 2000). Although a large number of compounds are marketed for inhibition of specific kinases, caution should be used when choosing an inhibitor and interpreting its effects on signaling. ATP binding sites are highly conserved across the kinome (Manning, Whyte, Martinez, Hunter, & Sudarsanam, 2002), and most inhibitors target multiple kinases, either within a family or in different branches of the kinome (Fabian et al., 2005). Researchers should familiarize themselves with these off-target effects and use the lowest effective concentration of inhibitor to disfavor weaker binding interactions. Furthermore, many kinase inhibitors are poorly soluble in aqueous buffers, necessitating formulation for experiments in vivo or pretreatment for experiments in vitro (Eckstein et al., 2014;Herbrink, Schellens, Beijnen, & Nuijen, 2016). A final consideration when working with ATP-mimetic inhibitors is that these inhibitors typically bind and may even induce the active conformation of the kinase. This can lead to a paradoxical increase in typical readouts of kinase activation (e.g., phosphorylation of the activation loop tyrosine) and may even ultimately promote signaling due to release of autoinhibition. Careful controls (e.g., phosphorylation of inhibitory/activating sites on the kinase and direct substrates) should be probed along with downstream readouts of cell activation. Ultimately, however, smallmolecule inhibitors for many kinases are well characterized and commercially available and require little up-front investment of time or resources. Moreover, a pharmacological approach can uniquely enable the study of transient effects with high kinetic fidelity and minimal regulatory compensation. Inhibitors are thus powerful tools for dissecting kinase contribution to immune activation.
Chemical-genetic methods for studying kinase signaling in immune cells combine the specificity of gene editing with the temporal control of small-molecule inhibitors. In one approach kinases are sensitized to a bulky analog of an ATP competitive kinase inhibitor by substituting a smaller amino acid side chain for the usual aliphatic, polar, or bulky gatekeeper residue (Lopez, Kliegman, & Shokat, 2014). Since the gatekeeper is not directly involved in ATP binding, the "analog-sensitive" kinase retains kinase activity until the designer inhibitor is added (Bishop et al., 2000). This chemical-genetic approach can be used in transfected cells or incorporated into the genome of a model animal as a transgene or knockin. Since endogenous kinases have more occlusive gatekeeper residues, the engineered kinase-inhibitor pair is much more specific than traditional kinase inhibition (Fig. 4). Importantly, analog-sensitive kinase inhibition has the additional advantage over genetic or siRNA knockout approaches in that normal kinase function in the absence of inhibitor will allow direct comparison of cells pre-and posttreatment. This real-time component also minimizes the likelihood of compensatory transcriptional changes and other adaptations in primary cells or animals and selective pressure and evolution in cell lines. This approach has been used to identify the specific roles for Zap-70 in T cell activation and Csk in T cell and macrophage activation, but the approach can be applied to other kinases as well (Freedman et al., 2015;Levin, Zhang, Kadlecek, Shokat, & Weiss, 2008;Tan et al., 2014). Furthermore, although many kinase inhibitors blunt signaling, some kinases such as Csk have paradoxical negative regulatory functions. Inhibition of Csk AS with 3-IB-PP1 leads to robust SFK activation (Freedman et al., 2015;Tan et al., 2014). Inhibition of these negative regulatory kinases can be used as potent stimuli of Brian et al.

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Current Protocols in Immunology cellular signaling and can be combined with other kinase inhibitors to tease apart kinase contribution to cellular activation and protein dynamics (Brian et al., 2019).
Although some information can be gleaned from unbiased total protein and panphosphotyrosine detection methods, immunoblotting is typically most effective when applied as a targeted, relatively low-throughput process, requiring antibodies raised against unique peptide sequences or sites of posttranslational modification. The best antibodies have minimal cross-reactivity with other molecules in the cell. Small-volume, higher-throughput apparatuses are available, but these systems are less amenable to combining antibodies and resolving multiple species in a single blot. Despite these caveats, immunoblotting remains a robust, sensitive, and adaptable technique (Kurien & Scofield, 2015). If epitope-specific antibodies are unavailable, immunoblotting can be combined with immunoprecipitation. For example, total protein immunoprecipitation can be followed with a pan-phosphotyrosine blot, and molecular weight can be used to infer the identity of phosphorylated protein (Freedman et al., 2015). Alternatively, cyanogen bromide fragmentation (Thofte et al., 2018) can resolve phosphorylation of individual sites on multiply phosphorylated proteins.
Freed of the requirement for specific antibodies, LC-MS/MS is an excellent exploratory technique for quantifying poorly studied proteins and sites of post-translational modification. This method is especially useful for multiply modified proteins that cannot easily be probed by blotting. Advances in LC-MS/MS have allowed researchers to quantify tyrosine phosphorylation via targeted and unbiased approaches (Dekker et al., 2018;Hu, Noble, & Wolf-Yadlin, 2016;Liu & Chance, 2014). Proteomics approaches use databases to identify enzyme-digested peptides following resolution by LC-MS/MS. Unbiased LC-MS/MS can identify novel sites of phosphorylation in a cell lysate but may miss low-abundance peptides. In contrast, targeted approaches using isotope-labeled reference peptides are highly sensitive and can be used to quantify abundance or novel sites of post-translational modification in cell lysates and in vitro kinase assays using recombinant or purified proteins.
Together, these protocols describe powerful tools for investigating tyrosine kinase and other cell modulatory signaling. The methods range from general and flexible when reagents are available (immunoblotting) to more focused (co-immunoprecipitation and LC-MS/MS). Together, they constitute a process for dissecting the kinetics and dynamics of signaling pathway activation, protein-protein interaction, and novel tyrosine phosphorylation that are essential for understanding how the many inputs that engage tyrosine kinases are involved in immune activation, allowing researchers to develop tools that modulate immune function by directing kinase signaling.

Critical Parameters
Basic Protocol 1: A million cells lysed in 400 μl lysis buffer should yield enough protein for analysis with near-infrared-conjugated secondary antibodies and an appropriate imager (e.g., LI-COR Odyssey). We have found that this ratio works well for macrophages, but the ratio may need to be adjusted (∼doubled) for smaller cells, such as primary T and B cells, Jurkat cells, and mast cells, depending on the protein being analyzed. It is possible to use fewer cells, but the lysis buffer volume should be scaled to maintain comparable protein concentrations. Before beginning, primary antibodies should be validated to ensure specificity to the desired protein being probed.
Basic Protocol 2: For each condition a large number of cells (8-40 × 10 6 ) is required to ensure immunoprecipitation of sufficient protein for subsequent analysis. Stringency of the buffer, incubation, and wash conditions should be optimized so that only specific, biologically relevant protein-protein interactions are preserved. Stringency of co-immunoprecipitation can be assessed by blotting for nonassociated and loosely associated proteins in immunoprecipitates. To increase the stringency of immunoprecipitation, researchers can screen different lysis detergents and increase the salt concentration (over the typical 150 mM) in the wash buffer. It is also critical to keep lysis buffers, wash buffers, and beads cold to prevent phosphatase and protease activity.
Basic Protocol 3: When working with gels prior to protease digest, it is critical to avoid keratin contamination. Be sure to wear gloves, a face mask, and a laboratory coat, working to limit breathing or leaning over the gel as much as possible. Surfaces, tools, and instruments should be thoroughly cleaned with tissue wipes and 70% ethanol prior to use. Iodoacetamide and DTT should be portioned and dissolved in appropriate buffers immediately before use to prevent degradation from light. Buffer conditions,