Protease-activated receptor-2 (PAR2) is a G-Protein Coupled Receptor (GPCR) activated by proteolytic cleavage to expose an attached, tethered ligand (SLIGRL). We evaluated the ability for lipid-tethered-peptidomimetics to activate PAR2 with in vitro physiological and Ca2+ signaling assays to determine minimal components necessary for potent, specific and full PAR2 activation. A known PAR2 activating compound containing a hexadecyl (Hdc) lipid via three polyethylene glycol (PEG) linkers (2at-LIGRL-PEG3-Hdc) provided a potent agonist starting point (physiological EC50 = 1.4 nM; 95% CI: 1.2–2.3 nM). In a set of truncated analogs, 2at-LIGR-PEG3-Hdc retained potency (EC50 = 2.1 nM; 1.3–3.4 nM) with improved selectivity for PAR2 over Mas1 related G-protein coupled receptor type C11, a GPCR that can be activated by the PAR2 peptide agonist, SLIGRL-NH2. 2at-LIG-PEG3-Hdc was the smallest full PAR2 agonist, albeit with a reduced EC50 (46 nM; 20–100 nM). 2at-LI-PEG3-Hdc retained specific activity for PAR2 with reduced EC50 (310 nM; 260–360 nM) but displayed partial PAR2 activation in both physiological and Ca2+ signaling assays. Further truncation (2at-L-PEG3-Hdc and 2at-PEG3-Hdc) eliminated in vitro activity. When used in vivo, full and partial PAR2 in vitro agonists evoked mechanical hypersensitivity at a 15 pmole dose while 2at-L-PEG3-Hdc lacked efficacy. Minimum peptidomimetic PAR2 agonists were developed with known heterocycle substitutes for Ser1 (isoxazole or aminothiazoyl) and cyclohexylalanine (Cha) as a substitute for Leu2. Both heterocycle-tetrapeptide and heterocycle-dipeptides displayed PAR2 specificity, however, only the heterocycle-tetrapeptides displayed full PAR2 agonism. Using the lipid-tethered-peptidomimetic approach we have developed novel structure activity relationships for PAR2 that allows for selective probing of PAR2 function across a broad range of physiological systems.
Citation: Boitano S, Hoffman J, Tillu DV, Asiedu MN, Zhang Z, Sherwood CL, et al. (2014) Development and Evaluation of Small Peptidomimetic Ligands to Protease-Activated Receptor-2 (PAR2) through the Use of Lipid Tethering. PLoS ONE 9(6): e99140. https://doi.org/10.1371/journal.pone.0099140
Editor: Roland Seifert, Medical School of Hannover, Germany
Received: March 17, 2014; Accepted: May 9, 2014; Published: June 13, 2014
Copyright: © 2014 Boitano et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All data are included within the manuscript.
Funding: This manuscript was developed through a multi-principal investigator collaboration between S.B., T.J.P. and J.V. This work was primarily supported by a multi-principal investigator grant from the National Institutes of Health (NIH)/NINDS, NS073664 (S.B., T.J.P., J.V.). This work was additionally supported by the following grants: NIH/NIEHS Superfund Research Grant ES04940 (S.B.; Raina Maier, P.I.); NIH/NINDS Grant NS065926 (T.J.P.); NIH/NIAID Grant AI083403 (S.B.; Michael O. Daines, P.I.); NIH/NIGMS Grant GM087369 (X.D.); NIH/NINDS Grant NS054791 (X.D.); and the Technology and Research Initiative Fund from Arizona State Proposition 301 (J.V.). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: T. P. is a PLOS ONE Editorial Board member. This does not alter the authors' adherence to PLOS ONE Editorial policies and criteria.
Protease-activated receptors (PARs) are a sub-family of G-protein coupled receptors (GPCRs) that have a unique mode of activation. PARs contain an embedded ligand that is exposed following proteolytic cleavage of the extracellular oriented NH2 terminus . The different N-termini of the PARs present substrates for a variety of proteases that create selective activation (or inactivation) mechanisms for signal transduction , , . The most common, diffusionally limited “tethered ligand” uncovered following trypsin-like serine protease activity of PAR2 [exposing SLIGKV (human) or SLIGRL (rodent)] serves as a potent agonist to the receptor. As an obvious consequence of its activation mechanism, PAR2 is associated with pathologies that have a strong protease release, including inflammatory related diseases such as arthritis, asthma, inflammatory bowel disease, sepsis, and pain disorders , , . Stimulation of PAR2 in pain-sensing primary sensory neurons (nociceptors) leads to the sensitization of a variety of receptors including the noxious heat and capsaicin receptor TRPV1 , , . This sensitization of sensory neuronal channels underlies thermal , ,  or mechanical hypersensitivity , ,  elicited by activation of PAR2. The involvement of PAR2 in pain and other pathologies makes it a prime target for drug discovery. Importantly, PAR2 has been associated with itch based partly on data obtained using the relatively potent PAR2 signaling peptide, SLIGRL-NH2. It is now clear that this peptide also stimulates an additional GPCR, Mas1 related G-protein coupled receptor type C11 (MrgprC11), and this receptor is responsible for the pruritic properties of SLIGRL-NH2 . Therefore, assessing the selectivity of PAR2 ligands against receptors that are selectively expressed in sensory ganglia (e.g., MrgprC11; , ) is critical to developing selective probes for PAR2.
Small peptides or peptidomimetics that mimic the ligand binding properties of the tethered ligand exposed by proteolysis of the N-terminus of the receptor have been used to directly activate PARs , , , . Activating peptides (e.g., SLIGKV-NH2 and SLIGRL-NH2) and peptidomimetics (e.g., 2-furoyl-LIGRLO-NH2  and 2at-LIGRL-NH2 ) have provided useful tools for establishment of structure-activity relationships (SAR) and rational drug design because they limit off-target effects that are often a complication of natural protease activation. Early SAR studies suggested that the minimal peptide sequence required for PAR2 activation is a pentamer (either SLIGR-NH2 or the less potent LIGRL-NH2 , ). More recently, heterocycle-dipeptide mimetics have been shown to retain PAR2 activity . However, full characterization of these shortened compounds has been hindered by a lack of assays sufficiently sensitive to evaluate full concentration responses. Commonly used assays require high concentrations (> 50 µM) that potentially limit PAR2-selectivity or prevent full solubility for preferred Ca2+ activation studies . It is now evident that a variety of GPCRs, including PAR2, can elicit signaling pathway-specific activation with distinct physiological responses , , , , , . A means to establish better evaluation of the minimal peptidomimetic structure required for full PAR2 activation would benefit PAR2 ligand discovery efforts.
Lipidation of peptide receptor agonists has been used to increase their potency via a variety of mechanisms . Because of the naturally tethered ligands in PAR2, we hypothesized that lipidation of peptide and peptidomimetic agonists could provide a membrane bound tether to better mimic the natural receptor activation and thus increase their potencies . Modification of the potent PAR2 peptidomimetic agonists 2at-LIGRL-NH2 and 2at-LIGRLO-NH2 with polyethylene glycol (PEG) spacers and a hexadecyl (Hdc) or a palmitoyl (Pam) group (e.g., 2at-LIGRL-PEG3-Hdc or 2at-LIGRLO(PEG3-Pam)-NH2) improves ligand potency to the low nanomolar range without sacrificing specificity to PAR2 as demonstrated in cell lines or in cells isolated from PAR2 wild type vs. PAR2-/- mice . Because of this increased potency, we hypothesized that this synthetic tethered ligand (STL) approach could be used to more closely examine SAR of peptidomimetics in an effort to better understand the minimal components necessary to specifically activate PAR2. In this report, we used the STL approach coupled with real time cell analysis (RTCA) and digital Ca2+ imaging microscopy to evaluate 14 compounds. We describe six STL compounds consisting of full or truncated parent peptidomimetic (2at-LIGRL-NH2) linked to three PEGs and one Hdc and evaluate their potencies, efficacies and specificities at PAR2, including screening against MrgprC11, to determine a minimal sequence necessary for specific activation of PAR2 in vitro and in vivo. Moreover, we used a parallel approach to fully evaluate potency of heterocycle di- and tetra-peptide mimetics using Ser1 and Leu2 substitutions known to activate PAR2 , . These findings identify a minimal structure required for specific full and/or partial activation of PAR2 and thus, further elucidate highly potent and specific probes to examine the function of this receptor in vitro and in vivo.
Materials and Methods
Nα-9H-fluoren-9-ylmethoxycarbonyl (Nα-Fmoc) protected amino acids, 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluoro-phosphate (HBTU), and N-hydroxybenzotriazole (HOBt) were purchased from SynPep (Dublin, CA) or from Novabiochem (San Diego, CA). Aldehyde (4-(4-formyl-3-methoxyphenoxy)butyrylaminomethyl) resin and Rink Amide resins were acquired from Novabiochem (San Diego, CA). N,N’-diisopropylcarbodiimide (DIC) and diisopropylethylamine (DIEA) were purchased from IRIS Biotech (Marktredwitz, Germany). A Ng-2,2,4,6,7-pentamethyl-dihydrobenzofuran-5-sulfonyl side chain protecting group was used for Arg. 2-aminothiazole-4-carboxylic acid and 5-isoxazole-carboxylic acid were obtained from Combi-Blocks (San Diego, CA); hexadecyl (Hdc-NH2) amine was obtained from Sigma-Aldrich. Fmoc-protected version of PEG (1-(9H-fluoren-9-yl)-3,19-dioxo-2,8,11,14,21-pentaoxa-4,18-diazatricosan-23-oic acid) was obtained from Novabiochem (San Diego, CA). Reagent grade solvents, reagents, and acetonitrile for High Performance Liquid Chromatography (HPLC) were acquired from VWR (West Chester, PA) or Aldrich-Sigma (Milwaukee, WI), and were used without further purification unless otherwise noted. Compounds were manually assembled using 5 mL plastic syringe reactors equipped with a frit, and a Domino manual synthesizer obtained from Torviq (Niles, MI). The C-18 Sep-Pak™ Vac RC cartridges for solid phase extraction were purchased from Waters (Milford, MA).
Truncated analogs of 2at-LIGRL-PEG3-Hdc 1–6 and compounds 11–14 were prepared as previously published by solid-phase synthesis as summarized in Figure 1 on 4-(4-formyl-3-methoxyphenoxy)butyrylaminomethyl resin (aldehyde resin; 0.9 mmol/g) using Fmoc/tBu synthetic strategy and standard DIC-HOBt and HBTU activations . Compounds 7–10 were prepared on Rink amide resin (0.67 mmol/g). The synthesis was performed in fritted syringes using a Domino manual synthesizer obtained from Torviq (Niles, MI). All compounds were fully deprotected and cleaved from the resin by treatment with 91% trifluoroacetic acid (TFA; 3% water, 3% triisopropylsilane, and 3% thioanisole). After ether extraction of scavengers, compounds were purified by reverse-phase HPLC and/or size-exclusion chromatography (Sephadex G-25, 0.1 M acetic acid) to >95% purity. Compounds were analyzed for purity by analytical HPLC and MS by Electrospray Ionization or Matrix Assisted Laser Desorption Ionization - Time of Flight (ESI, MALDI-TOF; see below).
Reductive alkylation: hexadecyl amine (5 equivalents), sodium cyanoborohydride (5 equivalents) in 5% acetic acid in DCM (0.25 M solution), overnight; ii a) Fmoc-PEG(6 equiv), DIC (3 equivalents) for first coupling b) Piperidine/DMF (1∶9) for Fmoc deprotection iii Fmoc/tBu synthesis continued as follows: a) Fmoc-aa-OH (3 equivalents) activated by HOBt (3 equivalents), DIC (3 equivalents), or HBTU (3 equivalents), DIEA (6 equivalents) in DMF; b) Piperidine/DMF (1∶9) for Fmoc deprotection; iv TFA-scavenger cocktail (91%), water (3%), triisopropylsilane (3%), and thioanisole (3%) for 4 hr.
Reductive alkylation (Figure 1, step i)
The aldehyde resin was swollen in dichloromethane (DCM) for 2 hr, washed with DCM and 5% acetic acid in DCM. A mixture of hexadecyl amine (Hdc-NH2) (5 equivalents), Sodium cyanoborohydride (5 equivalents) in 5% acetic acid in DCM (0.25 M solution) was injected into the syringe reactor. The reaction mixture was stirred overnight. The resin was washed with DCM, 5% acetic acid in DCM, N,N’-dimethylformamide (DMF), 10% diisopropylethylamine (DIEA) in DMF, DCM. A small sample of the secondary amine resin was protected by an Fmoc group (∼30 mg of resin was treated with 10 equivalents of Fmoc-Cl, 10 equiv DIEA in DCM). Resin loading was assessed spectrophometrically (UV at 301 nm; 0.49 mmol/g).
Solid Phase Synthesis (Figure 1, steps ii-iii)
The aliquot of secondary amide resin from the previous step (10 µmol) was swollen in DCM, washed with tetrahydrofuran-DCM, and the Fmoc-PEG was coupled via symmetrical anhydride (6 equiv of Nα-Fmoc-PEG and 3 equivalents of DIC in tetrahydrofuran-DCM) overnight. An on-resin test using Bromophenol Blue was used for qualitative and continuous monitoring of reaction progress. Fmoc group was removed with 10% piperidine in DMF (2 min + 20 min). The resin was washed with DMF (3X), DCM (3X), 0.2 M HOBt in DMF (2X), and finally with DMF (2X). Fmoc-PEG and following Fmoc-protected amino acid were coupled using pre-activated 0.3 M HOBt ester in DMF (3 equiv of Nα-Fmoc amino acid or Fmoc-PEG, 3 equivalents of HOBt and 3 equivalents of DIC) monitored by Bromophenol Blue test. To avoid deletion sequences and slower coupling rate in longer sequences, the double coupling was performed at all steps with 3 equivalents of amino acid or Fmoc-PEG, 3 equivalents of HBTU and 6 equivalents of DIEA in DMF. Wherever beads still tested Kaiser positive, a third coupling was performed using the symmetric anhydride method (2 equivalent of amino acid and 1 equivalents of DIC in DCM). Any unreacted NH2 groups on the resin thereafter were capped using an excess of 50% acetic anhydride in pyridine for 5 min. When the coupling reaction was finished, the resin was washed with DMF, and the same procedure was repeated for the next amino acid until all amino acids were coupled. 2-aminothiazole-4-carboxylic acid and 5-isoxazole-carboxylic acid were attached to the resin as symmetrical anhydride (6 equivalents of acid and 3 equivalents of DIC in DCM-DMF).
Cleavage of Ligand from the Resin (Figure 1, step iv)
A cleavage cocktail (10 mL per 1 g of resin) of TFA (91%), water (3%), triisopropylsilane (3%), and thioanisole (3%) was injected into the resin and stirred for 4 hr at room temperature. The crude ligand was isolated from the resin by filtration, the filtrate was reduced to low volume by evaporation using a stream of nitrogen, and the ligand was precipitated in ice-cold diethyl ether, washed several times with ether, dried, dissolved in water and lyophilized to give off-white solid powders that were stored at −20°C until purified. The crude compound was purified by size-exclusion chromatography and preparative HPLC.
The purity of products was checked by analytical Reverse Phase-HPLC using a Waters Alliance 2695 Separation Model with a Waters 2487 dual wavelength detector (220 and 280 nm) on a reverse phase column (Waters Symmetry C18, 4.6×75 mm, 3.5 µm). Compounds were eluted with a linear gradient of aqueous CH3CN/0.1% CF3CO2H at a flow rate of 1.0 mL/min. Purification of ligands was achieved on a Waters 600 HPLC using a reverse phase column (Vydac C18, 15–20 µm, 22×250 mm). Peptides were eluted with a linear gradient of CH3CN/0.1% CF3CO2H at a flow rate of 5.0 mL/min. Separation was monitored at 230 and 280 nm. Size exclusion chromatography was performed on a borosilicate glass column (2.6×250 mm, Sigma, St. Louis, MO) filled with medium sized Sephadex G-25 or G-10. The compounds were eluted with an isocratic flow of 1.0 M aqueous acetic acid. The pure compounds were dissolved in deionized water or dimethylsulfoxide at approximately 1 mM concentrations. Structures were characterized by ESI (Finnigan, Thermoquest Liquid Chromatography-Quadruplet ion trap instrument) or MALDI-TOF (Bruker Reflex-III) with α-cyanocinnamic acid as a matrix). For internal calibration an appropriate mixture of standard peptides was used with an average resolution of 8,000–9,000. High resolution mass measurements were carried out on a Bruker Ultraflex MALDI TOF-TOF and an Apex Qh Fourier Transformation-Ion Cyclotron Resonance (9.4 T) high resolution instrument.
16HBE14o- cells, a SV40 transformed human bronchial epithelial cell line , were obtained through the California Pacific Medical Center Research Institute (San Francisco, CA, USA). Cells were maintained and expanded as previously described . Briefly, cell lines were expanded in tissue culture flasks prior to transfer to 96 well E-plates (Roche) for experiments. Flasks and 96 well E-plates were coated initially with a matrix coating solution (88% Lechner and LaVeck basal medium, 10% bovine serum albumin (BSA; from 1 mg/ml stock), 1% bovine collagen type I (from 2.9 mg/ml stock), and 1% human fibronectin (from 1 mg/ml stock solution) and incubated for 2 hr at 37°C, after which the coating solution was removed and allowed to dry for at least 1 hr. 16HBE14o- cells were plated at a concentration of 1×105 cells/cm2 and grown in Eagle’s Minimal Essential Medium supplemented with 10% Fetal Bovine Serum (FBS), 2 mM glutamax, penicillin and streptomycin (growth medium) at 37°C in a 5% CO2 atmosphere. Growth medium was replaced every other day until the cells reached confluence (5–7 days). Cells were then transferred to 96 well E-plates for RTCA, or collagen/fibronectin/BSA coated glass coverslips for Ca2+ imaging experiments.
Primary mouse tracheal epithelial (MTE) cells were cultured as described . Briefly, mouse tracheas were removed, washed in phosphate-buffered saline for 5 min at room temperature, cut lengthwise, and transferred to collection medium [1∶1 mixture of Dulbecco’s Modified Eagle Medium (DMEM) and Ham’s F12 with 1% penicillin-streptomycin) at 37°C. Tracheae were then incubated at 37°C for 2 hr in dissociation medium (44 mM NaHCO3, 54 mM KCl, 110 mM NaCl, 0.9 mM NaH2PO4, 0.25 µM FeN3O9, 1 µM sodium pyruvate, and 42 µM phenol red, pH 7.5; supplemented with 1% penicillin-streptomycin and 1.4 mg/ml pronase). Enzymatic digestion was stopped by adding 20% FBS. Epithelial cells were gently scraped from the tracheas, centrifuged at 100 x g for 5 min at room temperature. Cell pellets were washed in base culture medium (1∶1 mixture of DMEM and Ham’s F12 with 1% penicillin-streptomycin and 5% FBS) and centrifuged at 100 x g for 5 min at room temperature. MTE cells were resuspended in full culture medium (1∶1 mixture of DMEM and Ham’s F-12, 1% penicillin-streptomycin, 5% FBS, 15 mM Hepes, 3.6 mM sodium bicarbonate, 4 mM L-glutamine, 10 µg/mL insulin, 5 µg/mL transferrin, 25 ng/mL epidermal growth factor, 30 µg/mL bovine pituitary extract) and transferred to collagen/fibronectin/BSA coated tissue culture flasks. Cells were fed every other day for one week until transferred to 96-well E-plates for RTCA experiments.
CHO cells were cultured in DMEM supplemented with 10% FBS and 1% penicillin and streptomycin at 37°C in a 5% CO2 atmosphere. One day before transfection, CHO cells were plated in a 60 mm cell culture dish at a concentration of 1×105 cells/cm2 and grown without antibiotics. An MrgprC11 cDNA in pcDNA3.1 vector was transfected into CHO cells using Lipofectamine™ 2000 (Invitrogen) prior to transfer to coverslips for experiments. Coverslips were coated with 0.1 mg/ml poly-D-lysine (from 2 mg/ml stock) and incubated for 1 hr at room temperature, after which the coating solution was removed and the coverslips were washed twice with double distilled water. CHO cells were seeded on coverslips at a concentration of 1×105 cells/cm2 6 hr after transfection. Transfected cells were incubated for 24 hr prior to Ca2+ imaging.
In vitro physiological response screening
16HBE14o- cells on E-plates in growth medium and in a 37°C, 5% CO2 incubator were monitored for the establishment for relative impedance overnight every 15 min using the xCELLigence™ Real Time Cell Analyzer (RTCA, Roche) , , . When cells reached baseline impedance the next day, and prior to the experiment, the RTCA was moved to room air and temperature where full growth medium was replaced with 100μL modified Hank’s Balanced Saline Solution (HBSS) pre-warmed to 37°C. The RTCA was then allowed to come to room temperature (45–60 min) prior to ligand addition. Each well was then supplemented with 100 µL HBSS containing appropriate ligands to measure concentration response ranges in quadruplicate. Additional wells were used for vehicle controls. Relative impedance in each well was monitored every 30 sec over 4 hr. Peak responses, defined as the maximal change in Normalized Cell Index, were used to define maximal response concentrations and physiological EC50s for each ligand.
The use of primary cultured MTE cells required different treatments and resulted in reduced overall signaling. Briefly, MTE cells were transferred to E-plates in minimal culture medium (100 µL/well of 1∶1 mixture of DMEM and Ham’s F-12, 1% penicillin-streptomycin, 3.6 mM sodium bicarbonate, 4 mM L-glutamine) and allowed to adhere for 4 hr. At that time each well was supplemented with 2x concentration of agonist in minimal culture medium. Relative impedance in each well was monitored every 30 sec for up to 2 hr.
In vitro Ca2+ Imaging
16HBE14o- cells or CHO cells were loaded with fura 2-acetomethoxyl ester (CalBiochem or Molecular Probes) for 30 min at room temperature. Cells were washed with HBSS and allowed to sit for at least 20 min prior to digital imaging. For activation and desensitization assay experiments using 16HBE14o- cells, [Ca2+]i was measured as previously described . Experiments consisted of 20 sec of recording of cells in HBSS to determine resting [Ca2+]i, followed by a 10 sec wash to introduce ligand and up to 10 min of recording for ligand washes required for desensitization experiments. Briefly, fura-2 fluorescence was observed on an Olympus IX70 microscope with a 40X oil immersion objective after alternating excitation between 340 and 380 nm by a 75 W Xenon lamp linked to a Delta Ram V illuminator (PTI) and a gel optic line. Intracellular Ca2+ concentration ([Ca2+]i) for each individual cell in the field of view was calculated by ratiometric analysis of fura-2 fluorescence using equations originally published in . Individual ratios were calculated every sec throughout the experiments. [Ca2+]i traces over time are averaged [Ca2+]i of all cells within a field of view (80–120) and are representative of at least 3 experiments. Cells were considered activated by the ligand if their resting [Ca2+]i was increased above a 200 nM threshold, a 2–4 fold increase above typical resting values. Percent activation graphs are determined from 3–6 experiments for each ligand concentration. Time to threshold in individual cells represent between 200–400 cells from at least 3 experiments at each ligand concentration. Ca2+ imaging of CHO cells was similar; each ligand concentration tested included at least 3 experiments and a minimum of 200 cells analyzed during each experiment.
In vivo mechanical sensitivity
Male ICR mice (Harlan) or PAR2-/- mice and their wild type littermates on a C57Bl/6 background weighing 25–30 grams were used for these studies. Animal protocols were approved by the Institutional Animal Care and Use Committee of The University of Arizona. Compounds were injected into the plantar surface of the hindpaw in a total volume of 25 µL using a 31-gauge needle. Compounds were diluted using sterile saline. Mechanical thresholds were determined using calibrated von Frey filaments (Stoelting Co, Wood Dale, IN) with the up-down method . The experimenter was always blinded to the treatment conditions and animals were randomized such that animals in a single experimental group were never all housed together.
All statistical analyses were evaluated with GraphPad software (San Diego, CA). Multivariate comparisons were done with a two-way ANOVA with Tukey's or Bonferroni multiple comparison post-test as appropriate for the individual experiment. Pair-wise comparisons were done with a two-tailed Student's t-test. A value of p<0.05 was used to establish a significant difference between samples. Data in Figures are graphed ± Standard Error of the Mean (SEM) unless otherwise noted.
Determination of a minimal peptidomimetic structure needed for PAR2 activation using synthetic tethered ligands (STLs)
Truncated analogs of 2at-LIGRL-PEG3-Hdc, compounds 1–6, were synthesized using standard Fmoc chemistry on aldehyde amino methyl resin as described in Figure 1 and in detail in Flynn, et al . Briefly, compounds assembled on the solid support were cleaved from the resin with TFA-scavenger cocktails and purified by Reverse Phase-HPLC and/or size-exclusion chromatography. All compounds gave > 95% analytical HPLC and expected MS (data not shown).
In vitro potency.
Compounds 1–6 were first evaluated for in vitro physiological response using the xCELLigence™ RTCA. RTCA measures physiological interactions between the cellular membrane and a surface substrate using underlying electrodes that register changes in impedance (reported as a Cell Index) over a prolonged time course in a non-invasive system and has been used to evaluate PAR2 agonist potency , , . Each compound was applied to the cells over an appropriate concentration range and Cell Index was monitored over a 4 hr experiment. Agonist activity was similar between compound 1, the full length STL (2at-LIGRL-PEG3-Hdc; previously published as Compound 12 in Flynn, et al. ), and an STL missing the C-terminal Leu6, compound 2 (2at-LIGR-PEG3-Hdc), in their respective physiological responses (Figure 2) and in their calculated EC50s (compound 1 EC50 = 1.7 nM, 95% CI: 1.2–2.3 nM; 2 EC50 = 2.10 nM, 95% CI: 1.3–3.4 nM; Figure 3). Similar to previous RTCA analyses of PAR2 agonists, maximal response concentrations (solid lines in Figure 2) displayed faster return to baseline, and supramaximal concentrations (dashed black lines) resulted in reduced peak responses and faster returns to baseline , . Compound 3 (2at-LIG-PEG3-Hdc), constructed without two amino acids from the C-terminus of compound 1, displayed a reduced response in the RTCA assay (Figure 2) and a reduced EC50 (46 nM, 95% CI: 20–100 nM; Figure 3). Compound 4 (2at-LI-PEG3-Hdc), missing 3 amino acids from C-terminus of compound 1, was further reduced in RTCA response (Figure 2) and potency (EC50 = 310 nM: 95% CI 260–360 nM; Figure 3). The maximal response concentration of compound 4 also failed to reach a peak response similar to known full agonists 2-furoyl-LIGRLO-NH2, 2at-LIGRL-NH2 ,  or compounds 1–3, consistent with partial agonism of PAR2. Further C-terminus truncation (compounds 5 (2at-L-PEG3-Hdc) and 6 (2at-PEG3-Hdc)) eliminated ligand activity as measured by RTCA (Figure 2). Comparison of concentration response curves for compounds 1–4 shows relative equal potency for compounds 1 and 2, and measurable loss of potency (right shifts) for compounds 3 and 4 (Figure 4).
Each panel (A–F) represents physiological response, measured using xCELLigence™ RTCA and expressed as a Normalized Cell Index over time following addition of STL. (A) Compound 1, 2at-LIGRL-PEG3-Hdc; (B) Compound 2, 2at-LIGR-PEG3-Hdc; (C) Compound 3, 2at-LIG-PEG3-Hdc; (D) Compound 4, 2at-LI-PEG3-Hdc; (E) Compound 5, 2at-L-PEG3-Hdc; (F) Compound 6, 2at-PEG3-Hdc. Concentrations for each experiment chosen to highlight supramaximal (dashed black traces), maximal (solid traces) and concentration dependent responses are shown at right of individual plots. Traces are averages from three or four experiments and are representative of experiments from at least two independent E-plates. Standard deviations have been removed to promote clarity. Systematic truncation of the parent peptidomimetic reduces agonist response starting with compound 3 until no activity remains in compounds 5 and 6. The reduced peak responses at supramaximal and maximal concentrations of compound 4 are suggestive of a partial agonist.
Compound number, name, structure and the in vitro physiological EC50 (RTCA) of each compound described in the manuscript are shown for comparison. #, compound number; Name, compound name; Structure, compound structure; RTCA, xCELLigence™ real time cell analysis; EC50, half maximal effective concentration; 95% CI, 95% confidence interval.
Concentration response curves were developed from in vitro physiological responses (RTCA) using the peak response within the 4 hr experiment. Compounds 1 (2at-LIGRL-PEG3-Hdc) and 2 (2at-LIGR-PEG3-Hdc) have roughly equivalent EC50s (see Figure 3), while further truncated compounds 3 (2at-LIG-PEG3-Hdc) and 4 (2at-LI-PEG3-Hdc) have higher EC50s (see Figure 3).
Specificity of PAR2 agonists.
Although the RTCA experiments using 16HBE14o- cells provide a highly sensitive physiological assay that encompasses various cell signaling responses to an agonist, it is inherently limited in detecting receptor specificity. To further evaluate specificity of known and novel STLs, we first compared RTCA responses to compounds 1–4 using primary cultured mouse tracheal epithelial (MTE) cells from wild type and PAR2-/- mice (; Figure 5). The effective Cell Index for peak concentration response for each compound 1–4 was reduced in MTE cultures when compared to the 16HBE14o- cells. More importantly, compounds 1–4 all required PAR2 expression to establish RTCA responses. Also similar to the 16HBE14o- RTCA traces, compound 4 displayed a reduced peak response in the PAR2-expressing MTE. To demonstrate signaling competence in both cultures, stimulation with the PAR2 independent agonist ATP resulted in similar RTCA responses in both wild type and PAR2-/- primary MTE cultures.
Each panel (A–E) compares physiological response, measured with xCELLigence™ RTCA as a Normalized Cell Index over time following addition of STL in MTE cells cultured from PAR2 expressing mice (PAR2 wt, red traces) or PAR2 null mice (PAR2-/-, blue traces). Concentrations for each experiment are shown with each STL agonist (compounds 1–4) and a PAR2-independent agonist, ATP. Traces are averages from three or four experiments and are representative of experiments from at least two independent E-plates. Standard deviations have been removed to promote clarity. PAR2 expression is required for response to compounds 1–4, but not for response to ATP.
Because [Ca2+]i changes are a primary outcome following PAR2 activation, we further tested for PAR2 specificity using a Ca2+ desensitization assay with the known PAR2 agonist, 2at-LIGRL-NH2 , . Using digital imaging microscopy, we first evaluated minimal ligand concentrations that would induce 90–100% activation in 16HBE14o- cells within a 5 min experiment (Figure 6). Sample traces of average [Ca2+]i changes plotted over time for compounds 1–4 and the parent peptidomimetic, 2at-LIGRL-NH2 are consistent with RTCA recordings. Compounds 1 and 2 were highly potent ligands (15 nM) whereas compound 3 (300 nM) required higher concentrations to elicit the full Ca2+ response, albeit with a significant delay in the time required to reach threshold [Ca2+]i when compared with compounds 1 and 2 (Figure 6F, G). Compound 4 required even higher concentrations (2 µM) to achieve threshold [Ca2+]i changes. Even at this heightened concentration compound 4 displayed a significant drop in average peak [Ca2+]i as well as lack of return to baseline [Ca2+]i within the 5 min experiment (Figure 6D).
The top four panels (A–D) display traces from a single experiment of average individual cell [Ca2+]i (± SEM) over time for 16HBE14o-cells exposed to PAR2 STL agonist compounds 1–4. Concentrations (Compound 1: 2at-LIGRL-PEG3-Hdc, 15 nM; 2: 2at-LIGR-PEG3-Hdc, 15 nM; 3: 2at-LIG-PEG3-Hdc, 200 nM; 4: 2at-LI-PEG3-Hdc, 2 µM) were chosen to reflect minimal agonist concentration necessary to result in 95% activation of 16HBE14o- cells (n≥3 for each compound). The known peptidomimetic full agonist, (E) 2at-LIGRL-NH2 (10 µM) is shown for comparison. Although all compounds displayed full Ca2+ activation over the 5 min experiment (F), compounds 3 and 4 displayed a slight delay in average time to peak Ca2+ response (G).
For desensitization studies, 16HBE14o- cells were first exposed to a high concentration of 2at-LIGRL-NH2 to effectively eliminate PAR2 based signaling prior to application of compounds 1–4. Thus, any increase in [Ca2+]i in response to compounds 1–4 would indicate a response that was not specific to PAR2. When 16HBE14o- cells were desensitized with 50 µM 2at-LIGRL-NH2, a second wash with 50 µM 2at-LIGRL-NH2 did not result in an increase of [Ca2+]i (Figure 7A–D). Subsequent treatment with any of the four compounds also did not result in measurable changes in [Ca2+]i. This loss of response was not caused by loss of Ca2+ signaling itself, as ATP remained an effective agonist following desensitization of PAR2. We further tested the agonist ability of these novel PAR2 agonists by using 10 fold the fully activating concentration for each compound to desensitize 16HBE14o- responses to 10 µM 2at-LIGRL-NH2 (Figure 7E–H). In these experiments, compounds 1–3 effectively desensitized 16HBE14o- cells, however, compound 4 could not fully eliminate the 2at-LIGRL-NH2-induced Ca2+ response.
The top four panels (A–D) display traces of the average change in [Ca2+]i. for all cells in the field of view plotted over time (10 min). In each panel, PAR2 desensitization with 50 µM 2at-LIGRL-NH2 prevented Ca2+ signaling by a second application of 2at-LIGRL-NH2 and subsequent addition of PAR2 STL agonists — Compound 1: 2at-LIGRL-PEG3-Hdc, 15 nM; 2: 2at-LIGR-PEG3-Hdc, 15 nM; 3: 2at-LIG-PEG3-Hdc, 200 nM; 4: 2at-LI-PEG3-Hdc, 2 µM. Subsequent application of 5 µM ATP in each experiment demonstrated that Ca2+ response was intact, and only PAR2 dependent pathways were desensitized. In the bottom four panels (E–H), 16HBE14o- cells were desensitized with the STL compounds 1–4 at 10 fold their full activation concentrations Compound 1: 2at-LIGRL-PEG3-Hdc, 150 nM; 2: 2at-LIGR-PEG3-Hdc, 150 nM; 3: 2at-LIG-PEG3-Hdc, 2 µM; 4: 2at-LI-PEG3-Hdc, 20 µM). Although compounds 1–3 were fully effective in desensitizing the cells to 10 µM 2at-LIGRL-NH2, desensitization by compound 4 was not complete. In each case, responses to 5 µM ATP remained fully in tact.
It has been demonstrated that MrgprC11 can be activated by the PAR2 peptide agonist SLIGRL-NH2, (EC50 = 10 µM) however, the Leu6-truncated peptide, SLIGR-NH2, lost MrgprC11 signaling capacity while retaining PAR2 activity . To examine PAR2/MrgprC11 selectivity, we evaluated Ca2+ responses in MrgprC11 transfected CHO cells  with the parent peptidomimetic, 2at-LIGR-NH2 and the most potent STL compounds (1, 2) from this study (Figure 8). When 2at-LIGRL-NH2 was applied to MrgprC11 transfected CHO cells at very high concentration (10 µM), a modest Ca2+ response was observed (10% of cells in the field of view); no response was observed at 1 µM. In contrast, 10 µM of the full length PAR2-STL, compound 1, induced a robust Ca2+ response (88 ± 19%) in the MrgprC11 transfected cells. This Ca2+ response decreased profoundly at concentrations of compound 1 tested at 100 times higher than the RTCA EC50 of ∼ 1 nM (20% response at 1 µM, 10% response at 100 nM, 2.5% response at 10 nM, 0% at 1 nM). In contrast, the potent PAR2 agonist compound 2 (2at-LIGR-PEG3-Hdc) induced limited Ca2+ responses in the MrgprC11 transfected cells even at the highest concentrations tested (20% at 10 µM, 15% at 1 µM, 0.5% at 100 nM). All three compounds tested displayed selectivity for PAR2 versus MrgprC11 with 2at-LIGRL-NH2 and compound 2, 2at-LIGR-PEG3-Hdc, displaying minimal MrgprC11 activity at concentrations up to 10 µM.
Compounds were applied to PAR2 deficient CHO cells transfected with MrgprC11 and evaluated for Ca2+ response. Compound 1: 2at-LIGRL-PEG3-Hdc was able to induce a full Ca2+ response at the highest concentration tested that was negligible concentrations typically used for PAR2 activation (e.g., 1–10 nM). Both the parent compound (2at-LIGRL-NH2) and the Leu6-truncated STL (compound 2: 2at-LIGR-PEG3-Hdc) displayed limited MrgprC11 activity. None of the compounds displayed activity in untransfected CHO cells (not shown). Each column represents three experiments, each with ∼200 cells.
In vivo efficacy.
Stimulation of PAR2 in vivo is known to promote mechanical sensitization reflected by a mechanical hypersensitivity response in the von Frey test , , . Compounds 1–5 were individually injected into the hindpaw of mice following evaluation of baseline mechanical sensitivity and mechanical thresholds were evaluated at 1 and 3 hr post-injection. Based on the EC50s of parent compounds , ,  we utilized a dose of 15 pmoles for experimentation. Consistent with in vitro findings, compounds 1–4 evoked mechanical hypersensitivity at 1 and 3 hr following injection (Figure 9A–D) whereas compound 5 lacked activity (Figure 9E). We did not note any itch response following injection of any tested compound. To determine specificity of these ligands, we tested the parent compound for mechanical hypersensitivity in wild type (WT, C57Bl/6 background) and PAR2-/- mice (C57Bl/6 background). Compound 1 (15 pmoles) stimulated mechanical hypersensitivity in WT mice but failed to do so in PAR2-/- mice (Figure 9F). Therefore, these compounds are specific agonists at PAR2 in vivo with the minimal peptide sequence in vivo matching the in vitro activity.
Compounds were injected into the plantar surface of the hindpaw at 15 pmoles and mechanical sensitivity was measured at 1 and 3 hr following injection. Compounds 1: 2at-LIGRL-PEG3-Hdc, 2: 2at-LIGR-PEG3-Hdc, 3: 2at-LIG-PEG3-Hdc and 4: 2at-LI-PEG3-Hdc evoked mechanical hypersensitivity whereas Compound 5 (E; 2at-L-PEG3-Hdc) was inactive. Compound 1 caused mechanical hypersensitivity in PAR2 WT mice but was inactive in PAR2-/- mice (F). * p<0.05.
Evaluation of novel peptidomimetics using STL and RTCA.
A previous report suggested that the heterocycle Ser1 substitute isoxazole (io) combined with the amino acids cyclohexylalanine (Cha) and Ile3 (e.g., 5io-Cha-I-NH2) was sufficient to specifically activate PAR2 . However, in that report the authors also noted that full responses of this compound were not available due its lack of solubility and the sensitivity of the chosen Ca2+ assay used to evaluate PAR2 activation. Based on the previous report and our above data showing equipotency between compounds 1 and 2, we first looked at tetrapeptide mimetics that included the aminothiazoyl or isoxazole heterocycle paired with a Cha-IGR amino acid sequence and created compounds 7: 2at-Cha-IGR-NH2 and 8: 5io-Cha-IGR-NH2. We followed these with the proposed minimal heterocycle-dipeptides: compounds 9: 2at-Cha-I-NH2 and 10: 5io-Cha-I-NH2 (Figure 3). To evaluate potency of these compounds, we first took advantage of the highly sensitive nature of the RTCA in vitro physiological response (Figure 10). Compounds 7 (EC50 = 490 nM, CI: 370–640 nM) and 8 (EC50 = 240 nM, 95% CI: 170–320 nM) displayed RTCA EC50 responses consistent with high activity heterocycle-tetrapeptides, with compound 8 showing the most potent responses recorded for tetra-, penta-, or hexa-peptide mimetics used in this assay (e.g., 2at-LIGRL-NH2 and 2-furoyl-LIGRLO-NH2; ). Compounds 9 and 10 displayed reduced potency RTCA responses (compound 9: 1.1 µM, 95% CI: 810 nM–1.6 µM; 10 EC50 = 870 nM, 95% CI: 750 nM–1.0 µM; Figure 3). The isoxazole heterocycle demonstrated slightly higher potency when compared with similar length peptidomimetics containing the aminothiazoyl Ser1 substitute. Interestingly, from this shortened peptide group with the Leu2 Cha substitution, only compound 8 consistently displayed a Normalized Cell Index response consistent with full PAR2 agonism in the RTCA assay (Figure 10).
Each of the top four panels (A–D) represents physiological response to agonist compounds as described for Figure 2. Concentrations for each experiment (at right of plots) show concentration responses that include supramaximal (black dashed lines) and maximal (solid line) responses. Compound 7: 2at-Cha-IGR-NH2, compound 8: 5io-Cha-IGR-NH2, compound 9: 2at-Cha-I-NH2, and compound 10: 5io-Cha-I-NH2 all display rapid RTCA responses. However, compounds 7, 9, and 10, all exhibit reduced peak Normalized Cell Index responses. Concentration response curves developed from RTCA using the peak response within the 4 hr experiment are shown in the bottom panel. Compounds 7–10 display activity consistent with previously described heterocycle-pentapeptides PAR2 agonists , . EC50s for each compound are shown in Figure 3.
To better characterize differences among these heterocycle-tetrapeptides and heterocycle-dipeptides, we constructed companion STLs with two polyethylene glycol groups and a hexadecyl group (i.e., PEG2-Hdc attached to the C-terminus) and tested them for in vitro physiological responses with RTCA (Figure 11, Figure 3). Compounds 11 (2at-Cha-IGR-PEG2-Hdc) and 12 (5io-Cha-IGR-PEG2-Hdc) displayed RTCA EC50s in the nM range (EC50 = 16 nM, 95% CI: 12–20 nM and EC50 = 6.8 nM, 95% CI: 4.1–11 nM, respectively). Similar to the pattern observed above, truncation to the heterocycle-dipeptide STL, compounds 13 (2at-Cha-I-PEG2-Hdc) and 14 (5io-Cha-I-PEG2-Hdc), resulted in less potent agonists (13: EC50 = 86 nM, 95% CI: 59–130 nM and 14: EC50 = 43 nM, 95% CI: 28–65 nM). Additionally, both compounds 13 and 14 displayed a delayed onset of response that was most prominent at submaximal concentrations. Further, compound 13 clearly did not attain peak Normalized Cell Index responses observed by other compounds in this group. A beneficial outcome of increased sensitivity using the STL construction and RTCA analysis was the separation of potency when comparing compounds that only differed in their respective heterocycle head group. Notably, when Cha was substituted for Leu2 the isoxazole containing compounds displayed an increased potency over the aminothiazoyl containing compounds in their peptidomimetic form (compounds 7–10; Figure 10E, Figure 3) that was only clearly separable when tested in their STL form (compounds 11–14; Figure 11E, Figure 3).
Each of the top four panels (A–D) represents physiological responses to agonist compounds as described for Figure 2. Concentrations for each experiment (at right of plots) show concentration responses that include supramaximal (black dashed lines) and maximal (solid line) responses. Compounds 11: 2at-Cha-IGR-PEG2-Hdc, and 12: 5io-Cha-IGR-PEG2-Hdc both display rapid and full RTCA responses. However, compounds 13: 2at-Cha-I-PEG2-Hdc and 14: 5io-Cha-I-PEG2-Hdc display delayed responses across concentration ranges that fall short of peak Normalized Cell Index typical for a full agonist. (E) Concentration response curves developed from RTCA using the peak response within the 4 hr experiment are shown in the bottom panel. EC50s for each compound are shown in Figure 3.
We further characterized compounds 11–14, using the Ca2+ signaling assays. Typical Ca2+ traces with average [Ca2+]i changes (85–110 cells) plotted over time for compounds 11–14 are shown (Figure 12A–D). Concentrations for each compound were established by their ability to elicit 80–100% activation of 16HBE14o- cells above threshold (Figure 12E), and were consistent with the RTCA data in that the heterocycle-tetrapeptide STL constructions required lower concentrations than the heterocycle-dipeptide STLs. However, only compound 12 (5io-Cha-IGR-PEG2-Hdc) elicited Ca2+ traces consistent with a full PAR2 agonist. Both of the heterocycle-Cha-I-PEG2-Hdc compounds (13 and 14) could not consistently activate >95% of the cells in the 5 min experiment (Figure 12E). Examination of Ca2+ signaling data showed that compounds 11, 13 and 14 all exhibited a significantly delayed time to Ca2+ threshold following ligand application, and the heterocycle-dipeptide compounds also exhibited a reduced peak [Ca2+]i change (Figure 12F–G).
The top four panels (A–D) display traces from a single experiment of average individual cell [Ca2+]i (± SEM) over time for 16HBE14o-cells exposed to PAR2 STL agonist compounds 11–14. (E) Concentrations (Compound 11: 2at-Cha-IGR-PEG2-Hdc, 200 nM; 12: 5io-Cha-IGR-PEG2-Hdc, 50 nM; 13: 2at-Cha-I-PEG2-Hdc, 600 nM; 14: 5io-Cha-I-PEG2-Hdc, 300 nM) were chosen to reflect minimal agonist concentration necessary to result in the maximal activation of 16HBE14o- cells (n≥3 for each compound). (F) Compounds 11, 13 and 14 all demonstrated significantly delayed responses to Ca2+ peak and (G) compounds 13 and 14 also displayed significantly reduced peak [Ca2+]i changes. Of this group, only compound 12 displayed Ca2+ signaling responses representative of full PAR2 agonism.
Compounds 11–14 were subjected to Ca2+ desensitization assays to test for specificity of response. In desensitization assays using 50 µM 2-at-LIGRL-NH2 as the specific PAR2 ligand to desensitize 16HBE14o- cells, none of compounds 11–14 induced significant Ca2+ signaling (Figure 13A–D), consistent with PAR2 specificity for each of these compounds. Also as above, compounds 11–14 were used at 10x maximal Ca2+ signaling response concentrations to assay their ability to desensitization PAR2 responses in 16HBE14o- cells to 10 µM 2at-LIGRL-NH2. Although compounds 11 and 12 were able to desensitize Ca2+ responses in these assays, compounds 13 and 14 did not completely desensitize Ca2+ responses to 10 µM 2at-LIGRL-NH2 (Figure 13E–H). Examination of average Ca2+ responses following application of high concentrations of compounds 13 and 14 suggested only partial activation of the 16HBE14o- cells (Figure 13I).
The top four panels (A–D) display traces of the average change in [Ca2+]i. for all cells in the field of view plotted over time (10 min). In each panel, PAR2 desensitization with 50 µM 2at-LIGRL-NH2 prevented Ca2+ signaling by a second application of 2at-LIGRL-NH2 and subsequent addition of PAR2 STL agonists — Compound 11: 2at-Cha-IGR-PEG2-Hdc, 200 nM; 12: 5io-Cha-LIG-PEG2-Hdc, 50 nM; 13: 2at-Cha-I-PEG2-Hdc, 600 nM; 14: 5io-Cha-I-PEG2-Hdc, 300 nM were monitored. Subsequent application of 5 µM ATP in each experiment demonstrated that Ca2+ response was intact, and only PAR2 dependent pathways were desensitized. In the bottom four panels (E–H) 16HBE14o- cells were desensitized with the STL compounds 1–4 at 10 fold their full activation concentrations — Compound 11: 2at-Cha-IGR-PEG2-Hdc, 2 µM; 12: 5io-Cha-LIG-PEG2-Hdc, 500 nM; 13: 2at-Cha-I-PEG2-Hdc, 6 µM; 14: 5io-Cha-I-PEG2-Hdc, 3 µM. Although compounds 11 and 12 were effective in desensitizing 16HBE14o- cells to 10 µM 2at-LIGRL-NH2, desensitization by compounds 13 and 14 was incomplete. In each case, responses to 5 µM ATP remained fully intact. (I) Further examination of Ca2+ responses in 16HBE14o- cells demonstrated an incomplete Ca2+ activation for compounds 13 and 14 persisted at the high agonist concentrations used to desensitize the cells. These data support PAR2 specificity for each compound, however the heterocycle-dipeptide STLs do not support full agonistic responses.
We have used a high sensitivity in vitro physiological assay combined with synthetic tethered-ligand (STL) approach to evaluate distinct protease-activated receptor-2 (PAR2) ligand structure activity relationships (SAR). First, using the RTCA physiological assay, we were able to present a minimal peptide sequence required for full and partial PAR2 activation and fully characterize EC50s for these truncated compounds. The use of this minimal peptide sequence both in vitro and in vivo opens new avenues for drug discovery and probing of physiological function at this receptor. Second, we were able to optimize SAR for PAR2 ligands with differing, high activity heterocycle (5-isoxazol and 2-aminothiazoyl) substitution of Ser1, paired with amino acid sequences naturally occurring in PAR2 or the previously used cyclohexylalanine (Cha) substitution for Leu2. Such discovery, which is facilitated by the STL approach, is ideal to evaluate otherwise minimally potent and/or questionably selective compounds for PAR2 and thus, provide a solid backbone for drug discovery. Finally, we provide detail on PAR2 specificity of these compounds, an important point considering the recently discovered pharmacological similarity between PAR2 and MrgprC11.
We first evaluated minimal peptide sequence analysis using successive truncation of a known activating peptidomimetic linked to a spaced lipid tether (e.g., ). Truncated analogs of 2at-LIGRL-PEG3-Hdc exhibited a descending trend of PAR2 activation with a minimal cut off at compound 4, 2at-LI-PEG3-Hdc. The heterocycle-dipeptide-STL maintained in vitro concentration responses (EC50 = 310 nM, 95% CI: 262–360 nM) equipotent with the commonly used heterocycle-pentapeptide and heterocycle-hexapeptide (e.g., 2at-LIGRL-NH2, RTCA EC50 = 310 nM, 95% CI: 240–400 nM; 2-furoyl-LIGRLO-NH2, RTCA EC50 = 240 nM, 95% CI: 190–290 nM; 6-aminonicotinyl-LIGRL-NH2, RTCA EC50 = 430 nM, 95% CI: 350–530 nM; ).
An alternative approach to assaying minimal peptide structures is the use of single Alanine substitutions (Ala-scan) in SLIGRL-NH2 and Ca2+ activation assays to assess PAR2 activation , , . Collectively, the Ala-scan studies revealed the importance of Ser1 and Leu2 for peptide-induced activation of PAR2 with full loss of activity when the Leu2 was substituted with Ala across all assays. The effects of Ser1 substitution with Ala, however, was dependent on the cellular assay with one group demonstrating near complete loss of activity in PAR2 expressing kNRK cells , another showing significant shift in activity in PAR2 expressing oocytes , and the third demonstrating only a slight loss of activity in transfected mouse embryonic fibroblasts . Other substitutions were again consistent across assays, where Ala substitutions of Ile3 and Arg5 decreased PAR2 activation while substitutions at Gly4 and Leu6 did not appreciably alter potency. When multiple Ala substitutions were made to activating peptides, it was shown that SLAAAA-NH2 could not activate Ca2+ signaling in kNRK cells . Through our STL-truncation approach coupled with the sensitive, in vitro physiological responses of RTCA, we found minimal changes in PAR2 activation between the parent compound 1 (2at-LIGRL-PEG3-Hdc) when Leu6 was removed (compound 2), successive reductions in potency following removal of Arg5-Leu6 (3) and Ile4-Arg5-Leu6 (4) and a complete loss of potency following removal of Ile3-Gly4-Arg5-Leu6 (5) or Leu2-Ile3-Gly4-Arg5-Leu6 (6). The minimal activating sequence both in vitro and in vivo required Leu2 and Ile3 in addition to the heterocycle substitute for Ser1 (compound 4, 2at-LI-PEG3-Hdc). Interestingly, when Ala substitutions were introduced into the receptor and activity uncovered by trypsin activation, the naturally tethered SLAAAA sequence was sufficient for PAR2 activation, albeit a less than full cellular response . This minimal activation could not be duplicated using the STL approach, where compound 5 (2at-L-PEG3-Hdc), was inactive both in vitro and in vivo. It is possible that loss of activity in 5, could be caused by absence of a peptide backbone or a lost interaction with the side chain of Ile3 that may be required in the absence of trypsin cleavage of the receptor. The importance of a peptide backbone is apparent when comparing RTCA activity from compounds 3 (2at-LIG-PEG3-Hdc) and 4 (2at-LI-PEG3-Hdc). The relatively high potency of compound 3 (EC50 = 46 nM) was achieved by retention of the Gly3, amino acid without any side chain. Compound 4, however, displayed significantly reduced potency in addition to a delay in time to peak and a reduction in peak Normalized Cell Index. Subsequent reductions in the ability for compound 4 to fully activate Ca2+ signaling suggest that activation by this minimal sequence results in only partial agonism of PAR2.
PAR2 activation is traditionally monitored by Ca2+ response following Gq activation and subsequent Ca2+ responses (e.g., , , , , ). However, it is well accepted that activation of PAR2 by native proteases or peptidomimetics can result in the recruitment of a variety of G-Proteins and multiple signaling pathways , . The RTCA approach used herein to screen PAR2 agonists relies on the cellular physiological response that is resultant of the various signaling pathways activated by the candidate drug . Response patterns to individual compounds are reflective of the signaling pathways activated and as such, have been used to classify GPCR ligands into subgroups . Compounds 1–4 tested in these studies displayed RTCA responses consistent with PAR2 drugs that elicit both Ca2+ and MAPK signaling , , and do not appear to invoke “biased signaling” via PAR2 , , , . Comparison of RTCA responses from primary cultured mouse tracheal epithelial (MTE) cells obtained from wild type or PAR2-/- mice successfully demonstrated the need for PAR2 expression to invoke physiological responses to these compounds. Traditional “desensitization” studies using Ca2+ signaling responses confirmed PAR2 specificity of truncated analogues. Extension of the traditional desensitization studies using high concentrations of the newly designed STLs as the agent to desensitize PAR2 to a known specific peptidomimetic agonist, 2at-LIGRL-NH2 allowed for further understanding of compound/PAR2 SAR. For example, the inability of compound 4 to fully desensitize cells at these heightened concentrations is in agreement with the RTCA results that suggest partial agonism by this selective PAR2 agonist.
The prototypical peptide activator for PAR2, SLIGRL-NH2, has recently been shown to contribute to the itch response via an alternative GPCR known to be expressed selectively in sensory neurons, MrgprC11 , . Although this receptor is not expressed in 16HBE14o- or MTE cells, and thus not a contributor to the in vitro results, activation of this GPCR in in vivo experiments could profoundly affect specificity in pain/itch pathways. Application of the parent STL (compound 1, 2at-LIGRL-PEG3-Hdc) to MrgprC11 transfected CHO cells resulted in a robust Ca2+ response, however, this required > 5,000-fold the RTCA EC50 concentration. Compound 2, with a truncated Leu6 resulted in limited activity at MrgprC11 at 10 µM and no activity at 1 µM. From these experiments we conclude that retention of the Arg5-Leu6 is preferred for MrgprC11 activation by our STL compounds. Concentrations required to activate Ca2+ responses in transfected MrgprC11 cells demonstrate at least several hundred fold selectivity for PAR2 over MrgprC11 by the STL compounds. Finally, the lack of response by 2at-LIGRL-NH2 at the EC50 concentration for SLIGRL-NH2 suggests that the Ser1 substitution confers selectivity for PAR2 over MrgprC11 in the absence of tethering. Therefore, the approach taken herein has identified highly potent and selective compounds that can be utilized to selectively probe the function of PAR2 in sensory biology.
A previous study reported on PAR2 activation using peptidomimetic derivatives of the first three amino acids of the natural tethered ligand for PAR2 (e.g., Ser1-Leu2-Ile3-NH2) at relatively high concentrations (50 µM) and demonstrated partial PAR2 activation using Ca2+ signaling assays in HEK293 cells . Significantly, one compound from this group, 5io-Cha-Ile-NH2 (published as compound 9 in  and compound 10 in this report) had an estimated EC50 similar to the peptide activator SLIGRLI-NH2 . However, the authors noted that lack of solubility of 5io-Cha-I-NH2 at high concentrations (100 µM) prevented full EC50 determination in their assay. Based on the minimal differences in potency observed in compounds 1 and 2 above, we took advantage of the sensitivity of the RTCA and STL approach to better evaluate EC50s of the heterocycle-dipeptides along with longer heterocycle-tetrapeptides. This new group included Ser1 substitute heterocycles 2-aminothiazoyl and 5-isoxazol with the Leu2 substitute cyclohexylalanine (Cha) and in combination with Ile3 or Ile3-Gly4-Arg5 terminated with an amino group. We found that compounds 7–10 all elicited RTCA responses in 16HBE14o- cells, however, only compound 8 (5io-Cha-IGR-NH2) elicited a traditional rapid and robust RTCA response typical of full and specific PAR2 agonists (e.g., compounds 1–3 herein; , ). Although not as potent as compounds 1 and 2 above, heterocycle-tetrapeptide STLs were highly potent activators of 16HBE14o- cells, with RTCA EC50s of 16 nM (11) and 6.8 nM (12), and significantly more potent than their corresponding heterocycle-dipeptide STLs (13: EC50 = 86 nM; 14: EC50 = 43 nM). Direct comparison of 5-isoxazoyl heterocycle with 2-aminothiazoyl heterocycle substitutions resulted in an ∼2 fold decrease in RTCA EC50s in both the heterocycle-tetrapeptide and heterocycle-dipeptide STLs. Substitution of Leu2 with Cha reduced potency in the heterocycle-tetrapeptides (compare compounds 2 and 11), whereas the same substitution increased potency in the heterocycle-dipeptide construct (compare compounds 4 and 13). Although these latter comparisons are tempered by differences in PEG2 (compounds 9–12) vs. PEG3 spacers (compounds 1–4), such spacer differences using 2at-LIGRL- and 2at-LIGRLO- as parent groups in STLs did not alter potency across assays , and thus, the different PEG spacers likely do not alter these conclusions. Ca2+ desensitization assays using 16HBE14o- cells confirmed specificity of the compounds 11–14 for PAR2. However, high concentrations of compounds 13 and 14 could not fully activate Ca2+ response nor were they effective at desensitizing 16HBE14o- cells from activation by 10 µM 2-at-LIGRL-NH2. These data suggest that the heterocycle-tetrapeptide STLs fully and specifically activate PAR2, whereas the heterocycle-dipeptide STLs are PAR2 specific, yet partial agonists.
The use of lipid tethering combined with RTCA and supplemented with traditional Ca2+ signaling analysis allowed for more robust and interpretable SAR for PAR2, including smaller structural nuances that provide an efficient vehicle for future drug development. A strength of this sensitive, tethered ligand approach is the ability to test peptidomimetic ligands in a form that better mimics the natural activation of protease-activated receptors that results in a significant increase in potency. For example, RTCA allowed for separation of potency of peptidomimetic compounds (e.g., 4.5 fold differences in RTCA EC50 ranging from 240 nM to 1.1 µM among compounds 7–10). The increased potency of STL derivatives also allowed for accurate Ca2+ signaling studies and confirmation of partial agonism without non-specific effects associated with using high concentrations of newly developed untethered ligands that can obscure SAR. It is interesting that the partial RTCA and Ca2+ signaling agonist compound 4 (2at-LI-PEG3-Hdc) gave a similar response to the full agonists compounds in our in vivo assays. These data provide evidence that full agonists (or by analogy, full antagonists) to PAR2 may not be needed for full effects in vivo. It is accepted that our STL approach increases hydrophobicity in the ligand. Although increased hydrophobicity has traditionally been considered as a negative for building drugs, more recently lipidation of peptides has been recognized as a viable avenue for drug discovery . In closing, we propose that the STL approach will continue to lead to the discovery of high potency peptidomimetics and small molecules as this technique can better identify contrasts between compounds and the resulting higher quality SAR will be enriched with otherwise undetectable structures which may contribute to high fidelity design.
We thank Dr. Andrea N. Flynn for discussion on development and execution of the experiments, Renata Patek for chemical synthesis expertise, Aubrey M. Cunningham for her help with the PAR2-/-mice, Daniel X. Sherwood for software that allowed for Ca2+ signaling comparisons and Terri Boitano for her copyediting contributions.
Conceived and designed the experiments: SB TJP JV XD. Performed the experiments: SB JH DVT MNA ZZ CLS YW JV. Analyzed the data: SB JH DVT MNA ZZ CLS XD TJP JV. Contributed reagents/materials/analysis tools: SB XD TJP JV. Contributed to the writing of the manuscript: SB JH CLS XD TJP JV.
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