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Learn more: PMC Disclaimer | PMC Copyright Notice Blood Adv . 2026 Feb 5;10(7):2351–2362. doi: 10.1182/bloodadvances.2025017522 Search in PMC Search in PubMed View in NLM Catalog Add to search Targeting PAR1 biased signaling with parmodulin reduces thromboinflammation and acute lung injury in sickle cell disease Nirupama Ramadas Nirupama Ramadas 1 Blood Research Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Nirupama Ramadas 1, 2 , Kailyn Lowder Kailyn Lowder 1 Blood Research Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Kailyn Lowder 1 , Joshua Dutton Joshua Dutton 1 Blood Research Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Joshua Dutton 1, 2 , Rebecca Claire Kazen Rebecca Claire Kazen 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Rebecca Claire Kazen 2 , Rani Sellers Rani Sellers 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Rani Sellers 2 , Jacob T DeRousse Jacob T DeRousse 3 Function Therapeutics, Inc, Milwaukee, WI Find articles by Jacob T DeRousse 3 , Christopher Dockendorff Christopher Dockendorff 3 Function Therapeutics, Inc, Milwaukee, WI Find articles by Christopher Dockendorff 3 , Erica Marie Sparkenbaugh Erica Marie Sparkenbaugh 1 Blood Research Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC Find articles by Erica Marie Sparkenbaugh 1, 2, ∗ Author information Article notes Copyright and License information 1 Blood Research Center, The University of North Carolina at Chapel Hill, Chapel Hill, NC 2 Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, Chapel Hill, NC 3 Function Therapeutics, Inc, Milwaukee, WI ∗ Correspondence: Erica Sparkenbaugh, Department of Pathology and Laboratory Medicine, The University of North Carolina at Chapel Hill, 116 Manning Dr, 8114 Mary Ellen Jones, Chapel Hill, NC 27599; [email protected] Received 2025 Jul 2; Accepted 2025 Dec 26; Collection date 2026 Apr 14. © 2026 American Society of Hematology. Published by Elsevier Inc. Licensed under Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International (CC BY-NC-ND 4.0), permitting only noncommercial, nonderivative use with attribution. All other rights reserved. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/). PMC Copyright notice PMCID: PMC13068854 PMID: 41632640 Key Points • Biasing PAR1 signaling with Parmodulin reduces thrombin generation, inflammation, and endothelial activation in SCD. • Parmodulin protects sickle mice from lethal acute chest syndrome, highlighting biased PAR1 modulation as a promising therapeutic strategy. Visual Abstract Open in a new tab Abstract Protease activated receptor 1 (PAR1) is expressed by numerous cell types, including endothelial cells. Thrombin cleaves PAR1 at Arg41 and activates proinflammatory and barrier disruptive signaling. Alternatively, PAR1 is cleaved at Arg46 by activated protein C (APC) that is bound to endothelial protein C receptor (ECPR), which induces anti-inflammatory and barrier protective signaling. In sickle cell disease (SCD), we showed that thrombin-PAR1 signaling contributes to vascular stasis and, more recently, that PAR1-R41–biased signaling enhances inflammation, whereas PAR1-R46 signaling reduces thrombo-inflammation. We hypothesized that ECPR-PAR1-R46–biased signaling protects sickle mice from thrombo-inflammation. To test this hypothesis, Townes sickle mice were treated with parmodulin (parmodulin 2 [PM2, aka ML161] or NRD-21) to promote protective, anti-inflammatory PAR1-biased signaling. We found that PM2 significantly attenuated thrombin generation, inflammation, and endothelial activation and protected sickle mice from a model of lethal acute chest syndrome. These results suggest that using PM2 to block thrombin-PAR1 signaling while inducing APC-like signaling can promote cytoprotective, anti-inflammatory effects in mouse models of SCD. Introduction Sickle cell disease (SCD) is the most inherited hemoglobinopathy, affecting ∼7 to 10 million people worldwide. 1 A single nucleotide mutation in the gene for β globin generates hemoglobin S, which leads to chronic hemolysis and transient episodes of vaso-occlusive crisis 2 and, subsequently, to a hypercoagulable state, vascular inflammation, and end-organ damage. 3 We and others have shown that chronic activation of coagulation is a hallmark of SCD and is a central mediator that drives the disease pathology. 4 , 5 , 6 , 7 In addition to their role in thrombosis, coagulation proteases, such as factor X and thrombin, mediate proinflammatory signaling via the cleavage of protease activated receptors (PARs). PARs comprise a family of G protein-coupled receptors, PAR1-4, that are expressed on a variety of cell types across tissues. PARs are activated by proteolytic cleavage of their N-terminal exodomain, thereby generating tethered ligands that initiate classical G protein-coupled receptor signaling pathways. 8 , 9 , 10 PAR1 was first identified as the thrombin receptor on human platelets that promotes platelet activation and aggregation. 11 , 12 , 13 Mouse platelets do not express PAR1; thrombin interacts with murine PAR3 via its hirudin-like domain, which brings thrombin into proximity with PAR4, thereby facilitating its cleavage and activation. 14 In both human and mouse endothelial cells (ECs), PAR1 exhibits biased signaling, which depends on the specific protease, site of cleavage, and other coreceptors present. Canonically, thrombin cleaves PAR1 at Arg41 (R41), which leads to G protein-coupled proinflammatory and barrier disruptive signaling. 12 When bound to its surface receptor thrombomodulin, thrombin does not induce PAR1-R41 proinflammatory responses and instead converts protein C (PC) to activated PC (APC). 15 , 16 PC/APC binds to the endothelial PC receptor (EPCR), an essential cofactor that enhances APC activation. 15 Moreover, EPCR, thrombomodulin, and PAR1 exist in a lipid raft microenvironment that enables EPCR-bound APC to cleave PAR1 at Arg46 (R46), thereby recruiting β-arrestin 2 and leading to anti-inflammatory and barrier protective signaling through cross talk with the sphingosine-1-phosphate receptor 1. 10 , 17 , 18 , 19 Previously, we demonstrated that thrombin-PAR1 signaling contributes to vascular stasis in a mouse model of SCD, likely by stimulating the release of Weibel-Palade bodies, which leads to the presence of the adhesion molecules P-selectin (P-sel) and von Willebrand Factor (VWF) on the EC surface. 20 More recently, we investigated the distinct contributions of thrombin- and APC-dependent cleavage of PAR1 to the vascular complications of SCD. We generated SCD mouse bone marrow chimeras (SS BM ) using mice that harbor point mutations in PAR1, which render them insensitive to either thrombin (PAR1 R 41Q ) or APC (PAR1 R 46Q ). 21 We found that SS BM /PAR1 R41Q mice had reduced thrombin generation and inflammation, whereas SS BM /PAR1 R46Q mice had enhanced inflammation when compared with SS BM mice with normal PAR1 expression. 22 These data suggest that when canonical thrombin-PAR1 signaling is blocked (SS BM /PAR1 R41Q ), thrombo-inflammation and vascular congestion are limited. Therefore, we hypothesized that blocking thrombin-PAR1 signaling while inducing cytoprotective PAR1 signaling will decrease the vascular pathology of SCD. There are several strategies to target PAR1 signaling therapeutically. Recombinant APC or an engineered signaling-selective variant with 3 lysine-to-alanin mutations in amino acids 191-193 (3K3A-APC) represent other options that have been beneficial in rodent models of inflammatory diseases, such as sepsis, ischemia-reperfusion injury, diabetes, and wound healing. 23 , 24 , 25 , 26 Parmodulins (PMs) are a class of small-molecule allosteric modulators of PAR1 that are thought to reversibly bind to an intracellular site. PMs simultaneously block detrimental thrombin signaling while inducing cytoprotective pathways in a similar manner to APC. 27 , 28 Emerging evidence indicates that PMs reduce thrombosis 27 and inflammation 23 , 29 , 30 in mouse models of disease. The goal of this study was to investigate if pharmacologic activation of cytoprotective and anti-inflammatory PAR1-biased signaling via PM improves vascular inflammation in SCD. Methods Mice Four-month-old Townes SCD mice, which express human α-globin and either human sickle β-globin (β S β S , SS) or normal adult β-globin (β A β A , AA, or wild-type control) were used for experiments. They were bred in-house from heterozygous β A β S (AS) breeding pairs purchased from Jackson Laboratories. Male and female mice were used in equal proportions. All end points were analyzed for sex-specific differences; no significant sex-based effects were observed. Mice were housed in autoclaved, ventilated cages (Techniplast Greenline) and maintained on a 12-hour light:dark cycle with ad libitum access to autoclaved food and reverse osmosis water. Reagent preparation Recombinant mouse tumor necrosis factor-α (TNF-α; R&D Systems) was prepared in sterile saline at 0.2 mg/mL. PM2 (PM2/ML161, MedChemExpress, Monmouth Junction, NJ) 31 was diluted to 10 mg/mL in dimethyl sulfoxide (DMSO) and vortexed until clear, followed by 1:1 dilution in polyethylene glycol 800 (PEG-800) and vortexing until clear. This solution was diluted to 2 mg/mL in sterile saline. The final vehicle formulation was 10% DMSO, 10% PEG-800, and 80% sterile saline. In addition to PM2, we used a second-generation PM, NRD-21 (provided by C. Dockendorff, Function Therapeutics, Milwaukee, WI), which was synthesized according to a published protocol. 32 NRD-21 solution was prepared in the same manner and vehicle as PM2. Sample collection At the end of the studies described hereafter, the mice were anesthetized with isoflurane (3% in 100% oxygen), and blood was collected from the inferior vena cava into tubes containing 3.8% sodium citrate. The kidneys and livers were collected and fixed in 10% neutral buffered formalin. The lungs were inflated in situ with 0.5 mL 10% formalin. The total blood cell counts were determined using a veterinary complete blood count analyzer (HT5, Heska). Plasma was collected from the blood via centrifugation at 4000 rpm for 15 minutes at room temperature and was stored at ˗80°C for cytokine and endothelial activation analysis, as described hereafter. PM2 treatment Study 1: steady state evaluation of PM2 treatment AA and SS mice received vehicle or PM2 (10 mg/kg, intraperitoneal [IP], 5 mL/kg body weight) on days 1, 3, 5, 7, and 8. The blood and organs were collected, as described previously, 1 hour after the last treatment. Study 2: PM2 treatment, followed by TNF-α challenge In a separate experiment, mice were treated with vehicle or PM2 (10 mg/kg IP) as described previously. On day 8, mice were challenged with TNF-α (2 μg/kg IP), followed by PM2 treatment after 30 minutes and 180 minutes. The blood and organs were collected 5 hours after the TNF-α challenge, as described previously. 33 Heme-induced acute chest syndrome (ACS) in sickle mice This experiment was conducted using only SS mice because AA control mice do not develop acute lung injury when treated with hemin (referred to as heme). 34 Hemin (Frontier Scientific) was prepared in 0.25 M NaOH at 25 mM, pH 8.0, and then filtered (0.22 μm syringe filter). The filtered hemin solution was diluted in sterile normal saline to 10 mM for final dosing. NRD-21 was prepared in the same manner as PM2 with a final vehicle formulation of 10% DMSO, 10% PEG-800, and 80% sterile saline. Female SS mice were treated with 10 mg/kg PM2 (IP), 10 mg/kg NRD-21 (IP), or vehicle 30 minutes before heme infusion. Some mice were treated once daily for 5 days via oral gavage with vorapaxar (105 μg/kg in 0.5% DMSO). After 30 minutes, the mice were treated with 140 μmol/kg heme (retro-orbital infusion) and monitored for acute lung injury for 120 minutes. Acute lung injury was scored as decreased spontaneous movement, decreased response to gentle stimuli, and labored breathing. Upon reaching the end point criteria, the mice were euthanized by cervical dislocation, and the lungs were inflated with 10% formalin and fixed for histologic analysis of the lung injury. They were stained with hematoxylin and eosin (H&E) and examined by veterinary pathologists who provided qualitative descriptions of the pathology. The images were acquired with an Olympus BX43 microscope, U-TV1Xc camera, and CellSens Software. Analysis of proinflammatory cytokines and endothelial markers The plasma concentrations of proinflammatory cytokines and endothelial markers were quantified using commercially available enzyme-linked immunosorbent assay kits, specifically for soluble EPCR (MyBioSource), thrombin-antithrombin complex (TAT; Siemens Healthcare Diagnostics), the proinflammatory cytokines interleukin-6 (IL-6; R&D Systems) and high mobility group box1 (HMGB1, Tecan), and the endothelial activation markers soluble vascular cell adhesion molecule 1 (sVCAM-1; R&D Systems), soluble P-sel (sP-sel, R&D Systems), and VWF (Abcam). We validated that the excess heme in SS plasma does not interfere with the immunoassays (data not shown). Protein isolation and western blot analysis Kidneys and livers were lysed using RIPA buffer (Cell Signaling Technology) containing 1% phosphatase-protease inhibitor cocktail (Sigma Aldrich). Kidney protein (70 μg) or liver protein (50 μg) were fractionated using sodium dodecyl sulfate–polyacrylamide gel electrophoresis with a Bio-Rad precision standard as molecular weight standards, followed by transfer onto polyvinylidene difluoride membranes (Bio-Rad Laboratories). The membranes were blocked using blocking buffer (Bio-Rad laboratories), followed by incubation with polyclonal antibody against EPCR (MyBiosource, MBS2003525,1:500 dilution). Beta-actin (MAB8929, R&D Systems; 1:3000 dilution for kidney, 1:10 6 for liver) was used to assess the total protein loading. Protein bands were visualized using horseradish peroxidase–conjugated secondary antibodies (Santa Cruz Biotechnology) and detected using enhanced chemiluminescence substrate (Bio-Rad Laboratories). Tissue histology The liver and kidney tissues that were collected after PM2 treatment were placed in 10% formalin for 24 hours and routinely processed using paraffin embedding. The samples were sectioned into 5-μm slices and stained with H&E. Liver congestion and necrosis were evaluated in the entire liver section using light microscopy at 20× magnification by 2 blinded researchers (N.R. and E.M.S.) in 5 nonoverlapping fields. Congestion was scored by the presence of sickled red cells in the sinusoids. A score of 0 indicated no red blood cells (RBCs) in the sinusoids; 1 indicated RBCs in <10% of sinusoids; 2 indicated RBCs in <25% of sinusoids; 3 indicated RBCs in <50% of sinusoids; 4 indicated RBCs in >50% of sinusoids, bridging between central veins, and RBCs present in extravascular parenchymal regions. Hepatic necrosis was identified by regions that lacked intact nucleated hepatocytes with eosinophilic inclusions and a lack of cytoplasmic membrane integrity. There were no consistent patterns of necrosis (eg, focal, zonal, centrilobular). A score of 0 indicated no obvious necrosis; 1 indicated necrosis in <10% of hepatocytes in the field; 2 indicated necrosis in <25% of hepatocytes in the field; 3 indicated necrosis in <50% of hepatocytes in the field; 4 indicated necrosis in >50% of hepatocytes and bridging between central veins. Hepatic fibrin(ogen) deposition was measured as described in the supplemental Methods . Kidney sections were additionally stained with periodic acid Schiff. Glomerulosclerosis, glomerular congestion, and interstitial fibrosis were qualitatively scored in a masked fashion by 2 veterinary pathologists (R.S. and R.C.K.) using the following scoring system: 0 indicated no finding; 1 indicated minimal finding; 2 indicated mild finding; 3 indicated moderate finding; 4 indicated marked finding; and 5 indicated severe finding. Statistical analysis Statistical analyses were performed using Prism 10 (version 10.3.1) software. The data were analyzed using either 1- or 2-way analysis of variance with Tukey’s post hoc test for multiple comparisons. For data that were not normally distributed, a Kruskal-Wallis test and Dunn’s multiple comparisons test were used. All experiments were approved by the University of North Carolina Institutional Animal Care and Use Committee. Results Increased EPCR shedding from renal vasculature and circulating EPCR suggest reduced APC-induced cytoprotection in sickle mice Endogenous APC binds to EPCR and activates PAR1, thereby inducing β-arrestin 2–dependent anti-inflammatory and cytoprotective signaling. 16 , 35 EPCR shedding has been observed in patients with SCD and mouse models and is linked to organ injury. 35 , 36 To investigate if there is increased shedding of EPCR in 4-month-old SS mice at steady state, the plasma levels of soluble EPCR (sEPCR) were quantified. There was a significant elevation in sEPCR in SS mice when compared with AA mice ( Figure 1 A). Furthermore, the EPCR levels in the SS kidneys were significantly reduced when compared with those of the AA controls ( Figure 1 B; supplemental Figure 1 ). We also assessed messenger RNA (mRNA) expression of EPCR in SS kidneys and observed no differences between the AA and SS tissues ( Figure 1 C). The liver is also a site of EPCR expression where it is found in the portal veins and venules. 37 Western blot and reverse transcription polymerase chain reaction analysis of EPCR protein and mRNA, respectively, revealed no differences in expression between the AA and SS tissues ( Figure 1 D-E). These data suggest that there was impaired endogenous APC-PAR1 signaling caused by shedding of EPCR in certain vascular beds in SS mice. Therefore, we used PM2 to investigate cytoprotective PAR1 signaling, bypassing the need for EPCR binding by APC or 3K3A-APC. Figure 1. Open in a new tab Loss of EPCR from the renal vasculature and increased sEPCR suggest loss of APC-induced cytoprotection in sickle mice. Plasma and kidneys were collected from 4-month-old AA (gray bars) and SS (blue bars) mice at steady state. (A) plasma levels of sEPCR (50% male). (B) Western blot analysis of total renal EPCR and β-actin expression. (C) mRNA analysis of renal EPCR and β-actin. (D) Western blot analysis of total hepatic EPCR and β-actin expression. (E) mRNA analysis of hepatic EPCR and β-actin. The data represent the mean ± standard error of the mean (SEM) of n = 4 to 5 replicates of the ratio of EPCR:β-actin. The data were analyzed using Student t tests. hprt, hypoxanthine phosphoribosyltransferase; ns, not significant. PMs protect against thrombo-inflammation and endothelial activation in sickle mice at steady state PMs simultaneously inhibit inflammatory thrombin-PAR1 signaling, mediated by Gq, while presumably promoting β-arrestin 2–dependent cytoprotective pathways in a similar manner as APC. 32 , 38 AA and SS mice were treated with vehicle or PM2 (10 mg/kg, IP) on alternating days for 1 week, and on the eighth day, samples were collected 1 hour after the last treatment ( Figure 2 A). PM2 had no effect on most of the hematologic parameters ( Table 1 ); however, it significantly reduced the levels of reticulocytes in SS mice. There was a significant elevation in TAT ( Figure 2 B), the inflammatory markers IL-6 and HMGB1 ( Figure 2 C-D), and the endothelial activation markers sVCAM1, VWF, and sP-Sel ( Figure 2 E-G) in SS mice when compared with the AA controls. PM treatment significantly decreased the elevated levels of TAT, IL-6, sVCAM, sP-sel, and VWF in SS mice at steady state. These results suggest that PMs can be used to preserve the cytoprotective signaling of PAR1 while blocking thrombin-dependent detrimental signaling. Figure 2. Open in a new tab PM inhibited thrombo-inflammation at steady state in sickle mice. Four-month-old AA and SS mice were treated with vehicle (gray bars) or 10 mg/kg PM2 (IP) (pink bars) on alternative days for 1 week, and samples were collected 1 hour after the last treatment (A). The plasma levels of (B) TAT, (C) IL-6, (D) HMGB1, (E) sVCAM, (F) VWF, and (G) sP-sel. The data are represented as the mean ± SEM, as analyzed by 2-way analysis of variance (ANOVA) and Tukey’s post hoc test. P values over brackets show the statistical comparisons. Table 1. Complete blood count analyses from AA, SS, and PM2-treated SS mice (n = 4-5 mice per group) Parameter AA AA+PM2 SS SS+PM2 RBCs, 10 6 /μL 10.61 ± 0.31 10.76 ± 0.22 6.14 ± 0.34∗∗∗ 6.44 ± 0.38∗∗∗ WBCs, 10 3 /μL 9.73 ± 0.87 6.89 ± 0.72 32.13 ± 4.08∗∗∗ 28.28 ± 2.78∗∗∗ Platelets, 10 3 /μL 1111 ± 59 1140 ± 55 907 ± 135 918 ± 58 Neutrophils, 10 3 /μL 1.84 ± 0.29 1.49 ± 0.18 7.18 ± 1.27∗∗ 5.8 ± 1.46∗∗ Monocytes, 10 3 /μL 0.31 ± 0.05 0.14 ± 0.02 0.87 ± 0.2∗ 0.42 ± 0.05 Hematocrit, % 34.9 ± 1.5 36.9 ± 0.9 29 ± 1.6 31.01 ± 1.5 MCHC, g/dL 30.3 ± 0.12 30.26 ± 0.17 27.07 ± 0.27 27.07 ± 0.21 Retic, % 7.58 ± 1.06 8.39 ± 0.36 57.33 ± 4.61∗∗∗ 35.49 ± 13.16∗∗ , † Open in a new tab The data are presented as the mean ± SEM (n = 9-15) as analyzed by 2-way ANOVA, followed by Tukey’s multiple comparison test. ∗ P < .05; ∗∗ P <0.01; ∗∗∗ P < .005 vs AA within the same treatment group. MCHC, mean corpuscular hemoglobin concentration. † P < .05 SS+PM2 vs SS. PMs modestly attenuate liver congestion and necrosis in sickle mice at steady state Because PM2 reduced biomarkers of thrombin generation, inflammation, and endothelial activation in SS mice at steady state, we investigated if PM2 had any effect on end-organ pathology. SS mice had elevated liver:body weight ratios that were not improved by PM2 treatment ( Figure 3 A). H&E liver sections were scored for vascular congestion and necrosis by 2 blinded observers (N.R. and E.S.). Sinusoidal congestion ( Figure 3 B) and necrosis ( Figure 3 C) were significantly elevated in SS mice, and both were modestly reduced by PM2 treatment. Representative H&E stained sections showed inflammatory infiltrates, congestion of sRBCs, and ischemic necrosis ( Figure 3 D). Although acute sickle hepatic crisis occurs in 10% of patients, which leads to transient increases in liver enzymes like alanine and aspartate aminotransferases, liver injury in SCD is primarily driven by chronic ischemia, inflammation, endothelial activation, and iron accumulation. 39 , 40 Therefore, we did not measure alanine aminotransferase or aspartate aminotransferase because these are not typically elevated in SS mice at steady state. 41 , 42 Figure 3. Open in a new tab PM attenuates hepatic congestion and fibrin(ogen) deposition. Quantification of the liver pathophysiology is presented as the congestion score (0-4; A) and the necrosis score (0-4; B) based on H&E-stained liver sections that were scored by 2 blinded observers. Each dot represents the average of 10 fields of view from 1 sample. Quantification of fibrin(ogen) deposition is represented as positive pixel count (C) using immunohistochemical staining of liver sections. (D) A representative image of an H&E stained section. Black boxes are magnifiied in the right panels. (E) Immunohistochemical stained section for fibrin(ogen) of liver tissues from AA and SS mice. Original magnification ×40; scale bars, 200 μm for H&E and 50 μm for fibrin(ogen) staining. The data are represented as the mean ± SEM, analyzed by 2-way ANOVA and Tukey’s post hoc test. ∗ P < .05; brackets represent comparison. C, congestion; i, inflammation; n, necrosos. The PM2-dependent reduction in TAT suggests reduced thrombin generation. To determine whether this translated into reduced hepatic fibrin(ogen) deposition, we performed immunohistochemical staining of fibrin(ogen) and quantified the positive pixels. Fibrin(ogen) deposition was modestly elevated in SS mice when compared with the AA controls, and PM2 treatment seemed to have no effect ( Figure 3 E). Fibrin(ogen) was predominantly localized around portal veins and within sinusoids, overlapping with regions of sRBC congestion ( Figure 3 F). We also investigated the effect of PM2 on kidney injury in SS mice. We and others have demonstrated that SS mice develop glomerulosclerosis, glomerular congestion, and tubular injury that progress with age. 43 , 44 Previous studies from our laboratory have shown that interventions that limit thrombin generation and inflammation can mitigate kidney injury in aged SS mice. 33 , 44 AA mice had no notable glomerular findings, and there was no statistically significant difference between the vehicle- and PM2-treated mice. The kidneys from SS mice were characterized by mild to moderate glomerulosclerosis ( Figure 4 A), mild glomerular congestion ( Figure 4 B), and minimal interstitial fibrosis ( Figure 4 C). SS mice had more Prussian blue staining in the proximal tubular ECs than AA mice, indicating more iron deposition ( Figure 4 E). PM2 treatment in sickle mice did not reduce fibrin and collagen deposition in the kidneys, as demonstrated by Martius Scarlet Blue (MSB) staining ( supplemental Figure 2 ). Overall, short-term PM2 treatment had no effect on the kidney pathology ( Figure 4 ). Figure 4. Open in a new tab PM has no effect on renal pathology. Quantification of glomerulosclerosis is presented as the sclerosis score (A) and congestion is presented as the glomerular congestion score (B) and interstitial fibrosis score (C) on a scale of 0 to 5, as scored by 2 blinded observers (R.S. and R.C.K.). (D) Representative image of an H&E stained section, and (E) representative images of Periodic acid-Schiff (PAS) stained sections of kidney tissue. Original magnification ×40; scale bars, 50 μm for both H&E and PAS staining. The data are presented as the mean ± SEM and were analyzed by 2-way ANOVA and Tukey’s post hoc test. There were no significant differences between the groups. Effect of PM2 on TNF-α challenge model of thrombo-inflammation We recently demonstrated that endogenous, noncanonical APC-PAR1-R46 signaling attenuates the heightened thrombo-inflammatory response in TNF-α–challenged SS mice. 22 To determine whether pharmacologic activation of noncanonical PAR1 signaling could replicate the protective effects of endogenous APC-PAR1-R46 signaling, AA and SS mice were treated with the biased PAR1 agonist PM2 every other day for 1 week. On the final day, the mice received TNF-α, followed by PM2 infusion at 30- and 180-minutes after the challenge and collection of the plasma 5 hours later ( Figure 5 A). As expected, TNF-α infusion markedly increased TAT, IL-6, and HMGB1 in SS mice ( Figure 5 B-D). However, PM2 treatment did not attenuate these responses. The endothelial activation markers (sVCAM-1, sP-sel, and VWF) were unchanged by the TNF-α challenge in SS mice and were not reduced by PM2 treatment ( Figure 5 E-G). Finally, TNF-α modestly increased sEPCR release in AA mice but did not exacerbate sEPCR shedding in SS mice ( Figure 5 H), suggesting that SCD itself drives sEPCR shedding to a threshold level. Collectively, these findings indicate that treatment with PM2 does not decrease measures of TNF-α–driven inflammation in wild-type or SS mice. Figure 5. Open in a new tab PM2 did not alleviate thrombo-inflammation in sickle mice after TNF-α challenge. Four-month-old AA and SS mice were treated with vehicle or PM2 (10 mg/kg IP) every other day for 1 week. On the eighth day, the mice were challenged with TNF-α (2 mg/kg IP), followed by a second dose of PM2 (10 mg/kg IP) 30 minutes later. Samples were collected 5 hours after the TNF-α treatment (A). The plasma levels of (B) TAT, (C) IL-6, (D) HMGB1, (E) sVCAM1, (F) sP-sel, (G) VWF, and (H) sEPCR were assessed. The data are presented as the mean ± SEM and were analyzed by 2-way ANOVA and Tukey’s post hoc test. The brackets indicate statistically significant comparisons. PMs protect sickle mice against heme-induced acute lung injury Thrombo-inflammation and endothelial activation are hypothesized to precipitate acute complications in SCD. ACS is one of the leading causes of hospitalization and mortality in patients with SCD and is linked to inflammation and endothelial activation. ACS has been modeled in SS mice with a high dose of hemin. 34 To investigate the role of biased PAR1 signaling, SS mice were treated with vehicle, PM2, 31 NRD-21, 32 or vorapaxar, an effectively irreversible orthosteric antagonist of PAR1, 45 , 46 30 minutes before infusion with a bolus of heme (140 μmol/kg, r.o.). The mice were monitored for signs of acute lung injury for 120 minutes. Heme treatment caused significant mortality (8/9 mice reached the end point) and a median survival time of 25 minutes. Pretreatment with vorapaxar produced similar results with a mortality rate of 4 of 6 and median survival time of 17 minutes ( Figure 6 A-B). Notably, both NRD-21 and PM2 protected the SS mice substantially from lethal acute lung injury ( Figure 6 A-B) with all NRD-21–treated mice surviving. Lung histopathology was also conducted. In a representative image from a vehicle-treated SS mouse that reached end point owing to heme-induced respiratory distress ( Figure 6 B), vascular congestion, alveolar wall thickening, and loss of architecture highlighted the acute lung injury. In contrast, the lung pathology was reduced in a PM2-treated mouse ( Figure 6 C). These results indicate that blocking all PAR1 signaling is not protective, whereas blocking canonical PAR1 signaling and promoting cytoprotective and anti-inflammatory signaling can protect against acute lung injury in SS mice. Figure 6. Open in a new tab PM2 and NRD-21 significantly protected SS mice from heme-induced ACS. SS mice received vehicle (red line) PM2 (10 mg/kg IP; pink line), NRD-21 (10 mg/kg IP; blue line), or vorapaxar (105 μg/kg, oral gavage; green line) 30 minutes before heme infusion (140 μmol/kg, retro-orbital). (A) Among the vehicle-treated mice, 3 of 16 mice reached the end point at a median survival time of 25 minutes. Among PM2-treated mice, 1 of 4 reached the end point. For NRD-21 treatment, 0 of 5 mice reached the end point. For vorapaxar, 4 of 6 mice reached the end point at a median survival time 17 minutes. The data show the survival proportions, as analyzed by the log-rank (Mantel-Cox) test. ∗ P < .05. (B) Representative image of a lung section from a heme-treated SS mouse that reached end point (t = 11 minutes). The lungs developed alveolar thickening (blue arrow), edema (green arrow), hemorrhage (red arrow), and vascular congestion (yellow arrow). (C) Representative image of a lung of a mouse that received PM2 before the heme challenge and that did not reach the end point. Images were taken at original magnification ×10. Discussion Despite the significant global burden of SCD, pharmacologic options are presently limited to hydroxyurea, L-glutamine, and crizanlizumab (with the approval of crizanlizumab withdrawn by the European Medicines Agency). The recent withdrawal of voxelotor from the market has left many patients and providers looking for other options. 47 Although the recent approval of transformative gene therapy has improved the disease outlook, these procedures require intensive clinical care and are currently limited to patients who are able to tolerate the chemotherapy required to ablate the immune system before the introduction of modified stem cells and costs $2 million to $3 million per patient. 48 Routine outpatient visits, unpredictable hospitalizations owing to complications, and medications accumulate to an average annual medical cost of $55 000 per patient with SCD in the United States. 49 Finally, these resources are further limited in the areas where SCD is most prevalent, namely Sub-Saharan Africa, the Indian subcontinent, and the Caribbean. There is clearly a critical need for affordable and accessible new therapeutic agents. In this study, we extended our recent findings that noncanonical APC-PAR1 signaling limits thrombo-inflammation in SS mice 22 by inhibiting thrombin-PAR1 signaling and promoting cytoprotective signaling with PM2. Treating SS mice with PM2 for 1 week significantly attenuated thrombin generation, inflammation, and endothelial activation at steady state. PM2 has been shown previously to reduce both factor Xa and thrombin generation on lipopolysaccharide- and TNF-α–stimulated HUVECs, as well as to decrease platelet adhesion to these activated ECs. 28 In addition, PMs are reported to suppress tissue factor (TF) expression on ECs in vitro. 32 We propose that PM2 attenuates TF exposure on the endothelium via endothelial barrier protection or endothelial repair, 27 , 28 thereby preventing blood contact with perivascular TF. In previous studies, we demonstrated that monocyte and perivascular TF are key drivers of thrombin generation in SS mice, 50 , 51 whereas endothelial TF drives inflammation. 52 These observations raise the possibility that PM2 may also reduce TF expression on monocytes. Furthermore, we also showed that the inhibition of TF or factor Xa also significantly lowers plasma IL-6 levels in SS mice at steady state. 52 The reduction in IL-6 observed in this study may similarly be a function of the indirect anticoagulant effects of PM2 or, alternatively, from direct anti-inflammatory signaling in ECs. Given that monocytes are also a prominent source of IL-6, it is also plausible that PM2 exerts additional effects on circulating white blood cells. Finally, the observed reductions in sVCAM-1, sP-selectin, and VWF likely reflect a direct cytoprotective effect of PM2 on ECs. PM2 also modestly reduced vascular congestion and hepatocellular necrosis in the livers of SS mice. It is not surprising that PM2 had no effect on the severity of light microscopic glomerular findings in SS mice, because 4-month-old mice already have substantial end-organ damage, and the duration of treatment was limited to 1 week. The study did demonstrate, however, that there were no obvious toxicities in the kidney at the dose evaluated. Although our previous data showed that preventing thrombin-PAR1 signaling in SS BM /PAR1 R41Q mice protected against the elevated thrombo-inflammation after TNF-α challenge, 22 PM2 did not produce a similar protective effect. One likely explanation is the short duration of treatment; SS mice received PM2 for only 1 week before the TNF-α challenge, whereas SS BM /PAR1 R41Q mice have lifelong inhibition of thrombin-PAR1 signaling. Supporting this interpretation, the only other intervention that demonstrated protection against TNF-α–induced thrombo-inflammation required long-term deficiency in coagulation factor XII. 33 Indeed, short-term inhibition of neither TF nor FXII was protective in this model. This suggests that sustained inhibition of detrimental signaling is necessary for therapeutic benefit in the context of the acute TNF-α challenge. In addition, PM2 is known to have a short half-life in mouse plasma, 31 and it is unable to inhibit thrombin-driven thrombus formation for a sustained period; however, its anti-inflammatory effects have been confirmed to be PAR1 dependent, 28 and APC and its variants also have a short half-life. 53 Future studies should explore whether APC variants with enhanced cytoprotective activity or combined approaches that inhibit thrombin-PAR1 signaling while promoting EPCR-dependent signaling offer greater therapeutic potential. Although TNF-α challenge is a validated and well-characterized model of vaso-occlusion used in the field, it causes both P-sel– and E-selectin–dependent adhesion of sickle RBCs and neutrophils to the activated endothelium and thus may not accurately replicate the clinical milieu of SCD. 54 , 55 Therefore, we investigated the role of PAR1 signaling in the pathology of a more clinically relevant acute stimulus, namely heme overdose. Indeed, we previously showed that PAR1 plays a role in heme-induced microvascular stasis through genetic PAR1 deficiency or inhibition with vorapaxar. 20 Unlike in our heme-induced microvascular stasis model, vorapaxar did not protect mice from heme-induced lethal acute lung injury. Importantly however, both PM2 and NRD-21 protected mice from lethal acute lung injury induced by a bolus of heme. Not only did SS mice survive the challenge, but they also had reduced markers of inflammatory lung injury based on histopathologic analyses. This supports the role of PMs in promoting cytoprotective PAR1 signaling and is consistent with their signaling bias when compared with vorapaxar, which inhibits both cytoprotective and proinflammatory PAR1 signaling. We also confirmed previously reported shedding of EPCR from the renal vasculature into the circulation. 36 In contrast, we did not observe differences in EPCR expression in the livers of SS mice. This is consistent with its limited distribution in this tissue where it is primarily located in the portal venous structures. Western blot analysis may lack the sensitivity to detect subtle changes in its expression; future studies will employ immunohistochemistry and confocal microscopy to more precisely evaluate EPCR expression in hepatic tissue. Analysis of EPCR mRNA expression in both tissues revealed no differences in the transcription levels between AA and SS mice, suggesting that EPCR protein is produced in SCD, yet subsequently from the tissue. However, the mechanism of EPCR shedding in SCD is unknown. EPCR can be enzymatically cleaved from the endothelium by proteases, such as TNF-α converting enzyme 56 , 57 and neutrophil proteinase 3. 58 These pathways will be investigated in future studies. Interestingly, PM2 had no effect on EPCR shedding at steady state or after TNF-α challenge, suggesting that reducing thrombo-inflammation alone is insufficient to stabilize EPCR expression. Limitations and future directions As previously noted, a limitation of our study is the short duration of treatment with PM2, the limited plasma stability of PM2, and the use of 4-month-old mice that have already accumulated significant vascular pathology. Future studies will investigate the effect of long-term treatment in preventing end-organ damage with treatment beginning in younger mice (eg, 1 month old) and lasting until 4 months. It is also important to consider the translational potential of this work. Because mice do not express PAR1 on platelets, our conclusions are limited to the effects of cytoprotective PAR1 signaling on ECs. PM2 prevents human platelet aggregation induced by thrombin. 27 , 31 Future studies using sickle human platelets will be critical to understand the hemostatic impact of PMs on platelets. Another option for targeting PAR1 in SCD is orthosteric PAR1 antagonists. Vorapaxar, for example, irreversibly binds the extracellular-ligand binding pocket, thereby preventing the activation of PAR1 by all proteases. 59 , 60 Vorapaxar is US Food and Drug Administration–approved for the treatment of myocardial infarction and peripheral arterial disease but carries a risk for intracranial hemorrhage when used as part of a dual antiplatelet regimen. 46 Patients with SCD have an elevated risk for stroke but are rarely prescribed dual antiplatelet therapy; however, vorapaxar use in patients with SCD is still unlikely. Vorapaxar has been shown to lack the protective effects of PMs on endothelium in vitro 27 and also in the ACS model described here. Because the platelet-inhibitory effects of PMs are reversible and less potent than vorapaxar, the risk for bleeding is likely reduced. Thus, it will be important to consider alternative approaches to interrupt thrombin-PAR1 signaling considering the therapeutic potential of APC-PAR1–biased agonism. Conclusion The understanding of SCD pathophysiology has significantly advanced in the last decade, but US Food and Drug Administration–approved treatments remain limited. Our findings demonstrate that inhibiting canonical thrombin-PAR1 signaling while promoting anti-inflammatory PAR1–biased signaling with a PM reduces thrombo-inflammation, decreases endothelial activation, and prevents lethal acute lung injury in a mouse model of SCD. This may be a promising and accessible approach for the treatment of SCD. Conflict-of-interest disclosure: C.D. reports being an owner and employee of Function Therapeutics, Inc, which is working to develop and commercialize parmodulins for future therapeutic applications. The remaining authors declare no competing financial interests. Acknowledgments E.M.S. is supported by National Heart, Lung, and Blood Institute (NHLBI) grants R43 HL169107 and R01 HL155193. C.D. reports funding from NHLBI grants R15HL127636 and R43HL169107. Authorship Contribution: N.R. performed the experiments, analyzed the data, and wrote the manuscript; J.D. and K.L. performed experiments; R.C.K. and R.S. performed the histopathologic analyses and wrote the manuscript; J.T.D. synthesized parmodulins; C.D. supervised the synthesis of parmodulins, assisted with the design of experiments, and edited the manuscript; and E.M.S. conceived of the study, designed and performed experiments, analyzed the data, and wrote and edited the manuscript. 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