Cardiac Targeting Peptide: From Identification to Validation to Mechanism of Transduction

Abstract

Cell-penetrating peptides (CPPs), also known as protein transduction domains, were first identified 25 years ago. They are small, ~6–30 amino acid long, synthetic, or naturally occurring peptides, able to carry a variety of cargoes across the cellular membranes in an intact, functional form. These cargoes can range from other small peptides, full-length proteins, nucleic acids including RNA and DNA, nanoparticles, and viral particles as well as radioisotopes and other fluorescent probes for imaging purposes. However, this ability to enter all cell types indiscriminately, and even cross the blood–brain barrier, hinders their development into viable vectors. Hence, researchers have adopted various strategies ranging from pH activatable cargoes to using phage display to identify tissue-specific CPPs. Use of this phage display strategy has led to an ever-expanding number of tissue-specific CPPs. Using phage display, we identified a 12-amino acid, non-naturally occurring peptide that targets the heart with peak uptake at 15 min after a peripheral intravenous injection, that we termed Cardiac Targeting Peptide (CTP). In this chapter, we use CTP as an example to describe techniques for validation of cell-specific transduction as well as provide details on a technology to identify binding partner(s) for these ever-increasing plethora of tissue-specific peptides. Given the myriad cargoes CTP can deliver, as well as rapid uptake after an intravenous injection, it can be applied to deliver radioisotopes, miRNA, siRNA, peptides, and proteins of therapeutic potential for acute cardiac conditions like myocardial infarction, where the window of opportunity for salvaging at-risk myocardium is limited to 6 hrs.

1. Introduction

The cell plasma membrane is a semi-permeable barrier that is essential for cell integrity and survival but at the same time presents a barrier to delivery of cargoes. Hence, the ability of trans-activator of transcription (Tat) protein of the human immunodeficiency virus to enter cultured cells and promote viral gene expression was met with great enthusiasm [1, 2]. Shortly thereafter, Antennapedia homeodomain, a homeobox transcription factor of Drosophila melanogaster, was shown to enter nerve cells and regulate neural morphogenesis [3]. Mapping of the domains responsible for this cell-penetrating ability led to the identification of two protein transduction domains, (also known as cell-penetrating peptides or CPPs); Tat corresponding to the 11 amino acid basic domain of HIV-1 Tat protein, and Penetratin corresponding to the 16 amino acid third helix of the Antennapedia domain. Subsequently, it was demonstrated that Tat fused to β-galactosidase and injected intraperitoneally in mice was internalized into multiple cell types including liver, heart, lung, kidney, and brain, delivering β-galactosidase in a functional form, highlighting the potential of Tat as a vector [4]. Currently, multiple cargoes in the form of peptides, proteins, nucleic acid, nanoparticles, and radioisotopes have been delivered using various CPPs [5, 6].

The ability of cationic or hydrophobic CPPs to transduce a wide variety of tissue types in vivo limits their utility because of lack of cell specificity. Phage display using libraries of various lengths and different bacteriophage strains have been utilized successfully to identify tissue-specific CPPs [7]. In our prior work, we identified a mildly basic, non-naturally occurring peptide (NH₂-APWHLSSQYSRT-COOH) capable of specifically targeting normal cardiomyocytes (CMCs) in vivo in mice, which we hence termed Cardiac Targeting Peptide or CTP [7, 8]. Our detailed bio-distribution studies show that peak uptake occurs at 15 min with complete disappearance of fluorescently labeled CTP by 6 hrs [9].

Cellular responses to ligands such as peptides, proteins, pharmaceutical drugs, or entire pathogens are generally mediated through interactions with specific proteins expressed on the cell surface. The ligand-based receptor capture (LRC)-TriCEPS methodology has been designed to directly identify such ligand–receptor interactions under near-physiological conditions on living cells. The key component of the LRC methodology is a trifunctional cross-linker that combines the following moieties; a N-hydroxysuccinimide (NHS), a hydrazone, and a biotin or an azide [10, 11].

In a typical LRC-TriCEPS experiment, at least two treatment arms are performed in parallel: one with the ligand of interest and a second with a control ligand with a known target. The first step of the LRC-TriCEPS experiment includes the conjugation of the ligand to TriCEPS using the NHS-ester. The TriCEPS-ligand conjugates are then incubated with previously oxidized cells. During this phase, the hydrazone captures covalently the surface glycoproteins via the aldehydes that are introduced at the carbohydrates by mild oxidation (receptor capture). After the receptor-capture reaction, the cells are lysed and the azide or the biotin groups are used to purify the cross-linked proteins for downstream mass spectrometry (MS)-based analysis (cell lysis, protein enrichment, and digestion). Upon identification, the cell surface proteins in the ligand samples are compared to those in the control sample using label-free quantification (Fig. 1). Randomly identified cell surface proteins are expected to have equal abundance in both samples, whereas the corresponding targets are found enriched in the ligand sample [10, 11].

Fig. 1.

Schematic of the workflow using LRC-TriCEPS technology. In a typical LRC-TriCEPS experiment, at least two treatment arms are performed in parallel: one with the ligand of interest and a second with a control ligand (i.e., a ligand with a known target). The first step of the LRC-TriCEPS experiment includes the conjugation of the two ligands to TriCEPS (ligand coupling). The TriCEPS-ligand conjugates are then incubated with previously oxidized cells under near-physiological conditions. During this phase, transient and stable ligand–receptor interactions will result in covalent capture events between TRICEPS and nearby carbohydrates (receptor capture). After the receptor-capture reaction, the cells are lysed, and cross-linked proteins are isolated and processed for mass spectrometry-based analysis (cell lysis, protein enrichment and digestion). Upon identification, the relative abundance of cell surface proteins in the ligand samples are compared to those in the control sample using label-free quantification. Randomly identified cell surface proteins are expected to have equal abundance in both samples, whereas the corresponding receptors are found enriched in the ligand sample

The LRC-TriCEPS methodology offers several unique advantages. It can be applied in a multitude of ligands ranging from small molecules to peptides, proteins, antibodies, and even whole viruses. Moreover, it does not require any genetic manipulation and therefore can be applied to a multitude of cell lines, including primary cells. Furthermore, targets are located within the context of the natural cell-specific surface microenvironment, so they are fully functional and exhibit their characteristic binding properties. Finally, it is hypothesis-free, meaning that no previous knowledge about the target is required.

A drawback of the methodology can be missing an interaction in case the target receptor is not expressed in the selected model system or TriCEPS coupling leads to hindering of the ligand–receptor interaction, which would interfere with ligand identification. In these scenarios, TriCEPS would not be useful. Additionally, the LRC-TriCEPS methodology enables the identification of only glycosylated targets; therefore, a small percentage of binding partners, less than 10%, cannot be identified.

In this chapter, we present details on the various methodologies that can be used to confirm tissue-specific internalization of a cell-specific CPP using CTP targeting the heart as an example. We also present details on LRC methodology.

2. Materials

2.1. Validation of Tissue-Specific CPPs

This section provides details on how to validate a candidate CPP for cell-specific transduction and contains protocols that can be modified for use in a variety of cell lines and organs, using fluorescently labeled peptides. We also provide a protocol for fluorophore conjugation of biotinylated CPPs. The methods detailed below use CTP in H9C2 cells, a rat cardiomyoblast cell line and wild-type mouse animal models as an example.

2.1.1. Transduction Assay Utilizing Fluorescence-Activated Cell Sorting or Confocal Imaging

1. Dulbecco’s Modified Eagle’s Medium—high glucose (DMEM).

2. Dulbecco’s Phosphate Buffered Saline (PBS).

3. Heat-inactivated fetal bovine serum (FBS).

4. Antibiotic-Antimycotic.

5. Rat Cardiomyoblast Cell Line (H9C2; ATCC).

6. Rat Cardiomyoblast Cell Line (H9C2 Cells) Media: Add 50 mL of heat-inactivated FBS and 5 mL of antibiotic-antimycotic to 500 ml of DMEM. Filter the mixture and store at 4 °C.

7. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA).

8. Immortal Mouse Fibroblast Cell Line (3T3; ATCC).

9. 16% Paraformaldehyde.

10. FACS Fixation Buffer: 10 mL of 16% PFA diluted with 30 mL of PBS. Solution should be stored with light protection at 4 °C.

11. Live/Dead Fixable Aqua Dead Cell Stain, 405 nm excitation. This dye is reconstituted in DMSO and is stored at −20 °C, light protected, when not in use.

12. Dimethyl Sulfoxide (DMSO).

13. Cy5.5-CTP-amide: Prepared by peptide synthesis, conjugated with Cy5.5 fluorophore, and purified using HPLC. Lyophilized powder is reconstituted in DMSO at 10 mM concentration. After reconstituting, store at −80 °C.

14. Glass-bottomed dishes (MatTek).

2.1.2. Bio-distribution In Vivo

1. 6- to 12-week-old albino, female mice (CD1, Charles River).

2. 28G 0.5 mL Insulin syringes.

3. 100 mg/mL Ketamine HCl (KetaVed).

4. 20 mg/mL Xylazine (AnaSed).

5. Ketamine/Xylazine Solution: Ketamine HCl is mixed with Xylazine 1:1 to produce a stock solution containing 50 mg/mL ketamine and 10 mg/mL Xylazine. This solution is made fresh before each use.

6. 26G Needles.

7. 3 mL Syringe.

8. 10% Buffered Formalin Phosphate.

9. Paraplast X-TRA (SIGMA).

10. Ethanol.

11. Xylene.

12. 10× Tris Buffered Saline (TBS).

13. 1× TBS Solution: 10× TBS is diluted 1:10 with deionized water. This solution can be stored at room temperature.

14. Microscope Slides.

15. Cover Glass.

16. Dapi Fluoromount G (SouthernBiotech).

17. Sparkle Optical Lens Cleaner.

18. Tissue-Tek Processing/Embedding Cassettes (ThermoFisher).

2.2. LRC-TriCEPS Methodology for Identifying Binding Partners

This section provides details on the materials needed to perform an LRC-TriCEPS experiment.

1. 5-methoxyanthranilic acid (Sigma-Aldrich).

2. Acetonitrile LC-MS grade (Fisher Scientific).

3. Alkyne agarose.

4. Ammonium bicarbonate.

5. Calcium chloride (CaCl₂).

6. Copper(II)sulfate (CuSO₄.

7. Ethylenediaminetetraacetic acid (EDTA).

8. Formic acid, 98–100% (vol/vol).

9. Glycine.

10. HEPES pH 8.2, 1 M buffer solution.

11. Iodoacetamide (Sigma-Aldrich).

12. Isopropanol for molecular biology, ≥99.5%.

13. Mobicol spin columns (Life Systems Design).

14. Mobicol filter 35 μm pore size (Life Systems Design).

15. PBS, pH 7.4.

16. Phosphoric acid 85 wt.% (H₃PO₄).

17. Protease inhibitors cOmplete (Sigma-Aldrich).

18. Rapigest SF (waters).

19. Sequencing grade modified trypsin (Promega).

20. Sodium ascorbate.

21. Sodium bicarbonate.

22. Sodium chloride.

23. Sodium dodecyl sulfate (SDS, Sigma-Aldrich).

24. Sodium hydroxide solution.

25. Sodium-metaperiodate (NaIO₄, Sigma-Aldrich).

26. TriCEPS v.3.0 (Dualsystems Biotech).

27. Tris (2-carboxyethyl)phosphine hydrochloride (TCEP, Sigma-Aldrich).

28. Tris (3-hydroxypropyltriazolylmethyl)amine (THPTA, Sigma-Aldrich).

29. Tris (hydroxymethyl)aminomethane base.

30. Urea.

31. Water HPLC gradient grade.

2.2.1. Buffer Preparation

1. Ligand coupling: HEPES 25 mM pH 8.2: Dilute 1 M HEPES with HPLC water and adjust the pH to 8.2 with NaOH.

2. Receptor capture: PBS pH 6.5: Acidify 500 mL of PBS (pH 7.4) by adding 105 μL of 85% (vol/vol) phosphoric acid.

3. Sodium (meta) periodate 1.5 mM: Dissolve 16 mg of sodium (meta) periodate in 50 mL of PBS pH 6.5 and vortex the solution until it is fully dissolved (always prepare fresh, protect from light).

4. 5-Methoxyanthranilic acid 5 mM: Dissolve 84 mg of 5-methoxyanthranilic acid in 100 mL PBS pH 7.4; stir with heating at 40 °C until it is fully dissolved and adjust pH to 7.4 with NaOH (protect from light).

5. Lysis buffer: Dissolve 4.8 g urea to 10 mL deionized water, stir until it is fully dissolved. Add 0.1% (vol/vol) Rapigest SF and one Protease Inhibitors cOmplete tablet, stir until it is fully dissolved (protect from light).

6. Protein enrichment: CuSO₄ 100 mM: Dissolve 25 mg of CuSO4 to 1 mL deionized water.

7. THPTA 100 mM: Dissolve 44 mg of THTPA to 1 mL deionized water (store aliquots in −20 °C).

8. Sodium ascorbate 1 M: Dissolve 200 mg of sodium ascorbate in 1 mL of deionized water (prepare fresh).

9. Click Chemistry Master Mix (2 ×): In 835 μL deionized water, add 125 μL THPTA, 20 μL CuSO4, and vortex; then add 20 μL sodium ascorbate and vortex again.

10. Tris-HCl 1 M: Dissolve 121.1 g of Tris base in 800 mL of deionized water; adjust the pH to 8.0 with HCl.

11. NaCl 5 M: Dissolve 292gm of NaCl in 1 L of deionized water.

12. EDTA 100 mM: Dissolve 1.86 g of EDTA to 50 mL of deionized water and adjust pH to 7.4 with NaOH.

13. SDS wash buffer (1% SDS, 100 mM Tris, 250 mM NaCl, 5 mM EDTA, pH 8): Mix 10 mL 1 M Tris–HCL, 5 mL NaCl, 5 mL 100 mM EDTA, and 1gm of SDS in a final volume of 100 mL.

14. Urea wash buffer (8 M urea in 100 mM Tris pH 8): Mix 48 g urea and 10 mL 1 M Tris to a final volume of 100 mL.

15. Isopropanol 80%: Mix 80 mL of isopropanol and 20 mL deionized water.

16. NaHCO₃ 100 mM: Dissolve 84 mg sodium bicarbonate in 100 mL deionized water, and adjust the pH to 11.0 with NaOH.

17. Acetonitrile 20%: Mix 20 mL of acetonitrile and 80 mL of deionized water.

18. Ammonium bicarbonate 50 mM: Dissolve 39 mg of ammonium bicarbonate in 10 mL of deionized water.

19. CaCl₂ 50 mM: Dissolve 55.5 mg in 10 mL deionized water.

20. Digestion buffer: Mix 10 mL 1 M Tris, 4 mL 50 mM CaCl₂, 10 mL acetonitrile to a total volume of 100 mL.

21. Formic acid 10%: Mix 10 mL of formic acid and 90 mL HPLC water.

22. Cl8 wash buffer: Mix 80 mL acetonitrile, 1 mL 10% Formic acid and 19 mL HPLC water.

23. Cl8 loading buffer: Mix 2 mL acetonitrile, 1 mL 10% formic acid, and 19 mL HPLC water.

24. Cl8 elution buffer: Combine 50 mL acetonitrile, 1 mL 10% formic acid, and 49 mL HPLC water.

3. Methods

3.1. Transduction Assay Using Fluorescence-Activated Cell Sorting

This protocol details how to perform an in vitro transduction assay of a candidate CPP for the purposes of validation. Here we provide a protocol to validate the specific transduction of Cy5.5-CTP into H9C2 cells with 3T3 mouse fibroblasts as a negative control.

1. Thaw cells from −180 °C liquid N₂ storage and plate onto a T-75 tissue culture treated flask with vented caps. Passage H9C2 cells for a minimum of three passages, trypsinizing at 60–70% confluence and plating at a 1:3 to 1:5 dilution (one 70% confluent T-75 flask yields ~one million cells) (See Note 1).

2. Passage the 3T3 cells for a minimum of 3 passages (similar to details given above for H9C2 cells), after thawing from −180 °C (see Note 1).

3. Once cells are 70% confluent, aspirate media, wash once with pre-warmed PBS, aspirate PBS and add 2 mL of Trypsin/EDTA, incubate for 5–7 min until cells have rounded up and are coming off in sheets.

4. Neutralize with 10–12 mL of pre-warmed media, collect cells, spin at 300 × g for 6 min to pellet the cells.

5. Aspirate supernatant and wash cell pellet twice with 10–12 mL of pre-warmed media.

6. Count cells using a hematocytometer and plate at a density of 1 × 10⁵ cells/well of a 12-well plate.

7. Twenty-four hours post-plating, aspirate and replace with pre-warmed 1 mL of media to which 10 μM of Cy5.5-CTP final concentration and 1 μL/mL of Live/Dead Fixable Aqua Dead Cell Stain has been added (see Notes 2 and 3). Return to incubator for 30 min. All cell works/groups/treatments are done in triplicate.

8. Suggested groups are H9C2/3T3 cells with no treatment or random peptide or CTP with Live/Dead stain added to all groups.

9. After the incubation period, wash cells extensively (at least 3 ×) with pre-warmed PBS, trypsinize, and centrifuge to collect cell pellet.

10. Wash the cell pellet with media once.

11. Aspirate the supernatant media above the cell pellet and resuspend in 1 mL of FACS Fixation Buffer.

12. Incubate the cells for 10 min at room temperature.

13. Add up to 5 mL of PBS and centrifuge to collect cell pellet.

14. Aspirate the supernatant and resuspend the cells in 200 μL of PBS.

15. Store the resuspended cells on ice, light protected.

16. In the flow cytometer run bleach and distilled water for 1 min each to clear the lines. Set to standby with a tube of distilled water.

17. Set up a protocol, specifying number of events to log and gates. Gates to set up would be to select for cells using FSC-A vs. SSC-A graph, a gate to select for live cells excluding the Live/Dead Fixable Aqua Dead Cell Stain using an SSC-A vs. 405 nm violet laser 535/50 filter, and graph to display SSC-A vs. 640 nm red laser 660/20 filter.

18. Vortex the cell sample, insert into the cytometer and adjust voltages so that the 660/20 fluorescence peaks in the middle of the graph. Apply these settings to the rest of the samples.

3.2. Transduction Assay Using Confocal Microscopy

This protocol is to acquire qualitative data about the transduction of a CPP using confocal microscopy. Here we utilize H9C2 cells incubated with Cy5.5 labeled CTP with 3T3 cells as a negative control. This protocol can be modified for use in ther cell lines, peptides, and fluorophores and can be modified for simultaneous fluorescent labeling of different cellular organelles for co-localization of CPP to particular cellular compartments.

1. Passage H9C2 cells and 3T3 cells as detailed in Subheading 3.1 above and plate in an optical glass-bottomed dish at a cell density of 5 × 10⁴ cells/well.

2. Twenty-four hours post-plating, aspirate, and replace with pre-warmed media to which Cy5.5-CTP has been added to give a final concentration of 10 μM.

3. Separate group of cells should be treated with vehicle (PBS) only and a random peptide similarly labeled with Cy5.5.

4. Return cells to incubator for 30 min. After the incubation period, wash cell extensively (at least 3 ×) with pre-warmed media. The cells are now ready to be imaged using confocal microscopy.

3.3. Bio-distribution In Vivo

This protocol details how to perform bio-distribution studies for a candidate CPP Here we provide an example of bio-distribution studies using Cy5.5-CTP in a mouse model. A number of organs should be harvested, such as the heart, lung, liver, kidney, brain, spleen, stomach, large intestine, small intestine, skeletal muscle, bone, and testes/ovaries, with each time point performed in triplicate.

1. Weigh mice and anesthetize with Ketamine/Xylazine (2 μL/g of tissue weight) administered intramuscularly or intraperitoneally. Adequate level of anesthesia will be achieved in 5–7 min, as assessed by lack of response to toe pinch.

2. Calculate a 10 mg/kg dose of Cy5.5-CTP, dilute to no more than 200 μL, and inject either retro-orbitally or through tail vein injection using an insulin syringe (see Note 2).

3. Allow peptide to circulate for the pre-specified time.

4. Euthanize mouse using Institutional Animal Care and Use Committee’s specified method and open the chest cavity.

5. Place a nick in the right atrium and, using a 26G needle, inject 3 mL of 10% Buffered Formalin Phosphate for the dual purpose of perfusion fixing the organs of the mouse and flushing out red blood cells.

6. Dissect out the organs of interest and store each organ individually in 10% Buffered Formalin Phosphate in a volume at least 20 times the volume of the tissue, weight per volume, for a minimum of 48 hrs, light protected, at room temperature (see Note 4).

7. Transfer organs into Tissue-Tek Processing/Embedding Cassettes and process the organs using a Tissue-Tek VIP processing machine.

8. Dehydrate the tissue in 70% EtOH for 30 min, followed by 80% EtOH for 30 min, 95% EtOH for 30 min, 95% EtOH for 30 min, 100% EtOH for 15 min, 100% EtOH for 20 min, and finally 100% EtOH for 20 min.

9. Clear the tissue in xylene twice, 30 min for each xylene treatment.

10. Infiltrate the cleared tissue with paraffin wax four times at 60 °C for 30 min with each treatment.

11. Embed in paraffin using metal molds. Fill the mold with molten paraffin kept at 65 °C and transfer to a cold plate. As the paraffin at the bottom of the mold begins to solidify, place the organ in the desired orientation. Place a labeled cassette on top of the mold as a backing and overfill with molten paraffin. Allow to cool until completely solid.

12. The block can then be stored in a −20 °C freezer over night to completely solidify.

13. Prepare a 38 °C water bath with distilled water.

14. Set up the microtome with a blade angle of 6° and a section thickness of 15 μm.

15. Section the tissue in the microtome by cutting the desired plane and then placing the blocks face down in the water back for 5 min or until the tissue has absorbed some moisture.

16. Place the tissue onto a flat ice block for 10 min.

17. Place a fresh blade onto the microtome and cut sections with a thickness of 8 μm. Discard bad paraffin ribbons until a ribbon of sufficient length and quality is produced.

18. Quality ribbons are then picked up with forceps and floated on the surface of the 38 °C water bath. Let the sections sit on the surface until they smooth out, taking care to not leave them too long to prevent the paraffin from disintegrating and tearing apart the section.

19. Float the flattened sections onto the surface of clean glass slides.

20. Place the slides into a 65 °C oven for 30 min to melt the wax. These slides can be stored at room temperature with light protection.

21. Deparaffinize the slides in Xylene three times, 10 min for each treatment.

22. Rehydrate the tissue in 100% EtOH for 5 min, followed by 95% EtOH for 5 min, 70% EtOH for 5 min, 50% EtOH for 5 min, and finally 1 × TBS for 5 min.

23. Mount the slides with coverslips using 125 μL of Dapi Fluoromount G.

24. Dry slides overnight at room temperature, light protected.

25. Image slides using confocal microscopy.

3.4. Ligand-Receptor Coupling Experimental Protocol for TriCEPS

This section provides details on how to perform an LRC-TriCEPS experiment and contains protocols that can be modified for use in a variety of cell lines and ligands of interest.

3.4.1. Ligand Coupling

The following protocol (Fig. 1) describes step-be-step an LRC experiment using TriCEPS v.3.0 (azide-containing) for generating one ligand and one control sample; therefore, the quantities need to be multiplied by the number of desired biological replicates (minimum three recommended) (see Notes 14-18).

1. Dissolve 100 μg of the ligand of interest in 50 μL of 25 mM HEPES (pH 8.2) in an Eppendorf tube (ligand sample).

2. Dissolve 100 μg of the control ligand in 50 μL of 25 mM HEPES (pH 8.2) in an Eppendorf tube (control sample).

3. If no control ligand is available, quench TRICEPS with 100 μg of glycine in 50 μL of 25 mM HEPES (pH 8.2).

4. Add 0.5 μL of the TRICEPS solution to each of the ligand and control sample and mix them thoroughly by pipetting.

5. Incubate the mixture at room temperature under constant gentle agitation for 90 min.

3.4.2. Receptor Capture (All Steps Performed at 4°C)

1. Collect 2 × 10⁷ cells with gentle scraping (for adherent cells) or centrifugation (for suspension cells) (see Notes 19 and 20).

2. Wash cells once with 50 mL PBS pH 6.5 in a 50 mL centrifuge tube.

3. Centrifuge the cells at 300 × g for 5 min at 4 °C, discard the supernatant, and resuspend the cells in the sodium (meta) periodate buffer.

4. Incubate cells at 4 °C in the dark for 15 min, under constant gentle agitation.

5. Centrifuge the cells at 300 × g for 5 min at 4 °C, discard the supernatant, and wash cells with 50 mL of PBS pH 7.4.

6. Centrifuge cells at 300 × g 5 min at 4 °C and resuspend cells in 20 mL 5-methoxyanthranilic acid buffer.

7. Split the cell suspension into two 15 mL centrifuge tubes.

8. Add the TRICEPS-coupled ligand of interest to one of the tubes and the coupled control ligand to the other.

9. Incubate ligands with cells for 90 min at 4 °C, under constant gentle agitation.

10. Centrifuge the cells at 300 × g 5 min at 4 °C and wash cells with 10 mL PBS pH 7.4.

11. Centrifuge the cells at 300 × g for 5 min at 4 °C, remove the supernatant, and freeze the cell pellets.

3.4.3. Cell Lysis and Protein Enrichment

1. Resuspend the cell pellets in 800 μL of lysis buffer and transfer the lysates to two Eppendorf tubes.

2. Sonicate the lysates using three 30 s sonication pulses and remove debris by centrifugation at 16,000 × g for 10 min.

3. Wash 200 μL alkyne agarose (per replicate) with 1.8 mL deionized water and add to the lysates.

4. Add 1 mL of 2 × click chemistry mastermix and incubate the samples for 18 h (±2 h) at room temperature under gentle agitation.

5. Pellet agarose beads by centrifugation for 4 min at 300 × g, remove supernatant, and wash beads with 1.8 mL deionized water.

6. Resuspend beads in 1 mL SDS wash buffer and reduce bead-bound proteins with 5 mM TCEP for 15 min at 55 °C and min at room temperature.

7. Pellet beads at 300 × g for 4 min to remove supernatant.

8. Alkylate the proteins with 40 mM iodoacetamide for 30 min, at room temperature, in the dark.

9. Transfer beads to Mobicol classic 35-μm filters and wash beads with 10 mL of the following buffers: SDS wash buffer, urea wash buffer, 5 M NaCl, 80% isopropanol, 100 mM NaHCO₃, 50 mM ammonium bicarbonate (60 °C), and 20% acetonitrile.

10. Transfer the beads in fresh Mobicol tube.

3.4.4. On-Bead Trypsin Digestion

1. Resuspend the beads in 400 μL digestion buffer.

2. Add 1 μg sequencing grade modified trypsin and incubate for 16 h at 37 °C.

3. Collect the peptides, wash beads twice with 50 mM ammonium bicarbonate.

4. Acidify samples with 10% formic acid to pH 3.

3.4.5. Peptide Purification

1. Desalt the peptides using UltraMicroSpin Cl8 Columns with 5–60 μg capacity for tryptic peptide fraction according to manufacturer’s instructions.

2. Dry the eluted peptides in a SpeedVac and store them at −80 °C until further analysis.

3.4.6. LC-MS/MS and Data Analysis

LRC samples are of medium-to-high complexity and need to be analyzed with a highly sensitive, high mass accuracy mass spectrometer. Analyze peptides with a standard shotgun mass spectrometry-based workflow and perform label-free quantification to extract relative protein abundance.

3.4.7. Statistical Analysis

Perform statistical analysis to calculate protein fold changes and their statistical significance between paired conditions; numerous free software packages exist for statistical analysis such as MSstats [12] and SafeQuant [13].

3.4.8. Data Visualization and Interpretation

A volcano plot combines a measure of statistical significance from a statistical test with the magnitude of the change, enabling quick visual identification of proteins that were significantly enriched in the ligand of interest samples. The x-axis represents the mean ratio fold change (on a log2 scale). The y-axis represents the statistical significance p-value of the ratio fold change for each protein (on a –log10 scale). Proteins that are enriched in one of the samples will plot either left or right of the x-axis origin, indicating in which sample the target protein is enriched.