Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2022 Jun 5.
Published in final edited form as: Biochem Biophys Res Commun. 2021 Apr 10;556:199–206. doi: 10.1016/j.bbrc.2021.03.161

Cell-free hemoglobin-mediated human lung microvascular endothelial barrier dysfunction is not mediated by cell death

Toria Tomasek a, Lorraine B Ware a,b, Julie A Bastarache a,b,c, Jamie E Meegan a
PMCID: PMC8483287  NIHMSID: NIHMS1692824  PMID: 33848934

Abstract

Circulating cell-free hemoglobin (CFH) contributes to endothelial injury in several inflammatory and hemolytic conditions. We and others have shown that CFH causes increased endothelial permeability, but the precise mechanisms of CFH-mediated endothelial barrier dysfunction are not fully understood. Based on our previous study in a mouse model of sepsis demonstrating that CFH increased apoptosis in the lung, we hypothesized that CFH causes endothelial barrier dysfunction through this cell death mechanism. We first confirmed that CFH causes human lung microvascular barrier dysfunction in vitro that can be prevented by the hemoglobin scavenger, haptoglobin. While CFH caused a small but significant decrease in cell viability measured by the membrane impermeable DNA dye Draq7 in human lung microvascular endothelial cells, CFH did not increase apoptosis as measured by TUNEL staining or western blot for cleaved caspase-3. Moreover, inhibitors of apoptosis (Z-VAD-FMK), necrosis (IM-54), necroptosis (necrostatin-1), ferroptosis (ferrostatin-1), or autophagy (3-methyladenine) did not prevent CFH-mediated endothelial barrier dysfunction. We conclude that although CFH may cause a modest decrease in cell viability over time, cell death does not contribute to CFH-mediated lung microvascular endothelial barrier dysfunction.

Keywords: endothelial, barrier dysfunction, hemoglobin, cell death

Introduction

Increased circulating levels of cell-free hemoglobin (CFH) occurring in hemolytic conditions such as sepsis, sickle-cell anemia, cardiac bypass surgery, trauma, pulmonary hypertension, and blood transfusion contribute to higher disease severity and organ damage [1, 2]. This is especially relevant to the lung, where these hemolytic conditions may cause disruption of the alveolar-capillary barrier leading to acute respiratory distress syndrome (ARDS), a condition characterized by accumulation of noncardiogenic pulmonary edema and severe hypoxemia [1, 3].

Normally, hemoglobin is safely confined within red blood cells, but under hemolytic conditions can be released into the circulation in large quantities [1, 4]. Red blood cells have highly effective antioxidant mechanisms that protect the cell from being exposed to high levels of reactive oxygen species (ROS) and keep the central iron atom in hemoglobin in a ferrous (2+) state [5]. When hemoglobin is released from the red blood cell, it is bound by its endogenous scavenger haptoglobin and the subsequent hemoglobin-haptoglobin complexes are cleared by macrophages. However, if this system is overwhelmed, free hemoglobin accumulates in the circulation where it is oxidized to the ferric (3+) state, having the potential to cause injury through a variety of mechanisms, including nitric oxide consumption, vasoconstriction, oxidative injury, activation of transcription factors, and alterations in cellular metabolism [1, 4].

We previously reported that CFH increased endothelial permeability and edema formation in the ex vivo isolated and perfused human lung and induced barrier dysfunction in human lung microvascular endothelial cells in vitro [6–8]. Furthermore, in a murine polymicrobial cecal slurry sepsis model, septic mice that received an additional injection of intravenous CFH had increased lung microvascular permeability and higher mortality compared to septic mice that received vehicle injections. Moreover, the septic mice injected with CFH had increased inflammation, oxidative stress, and endothelial cell apoptosis as measured by TUNEL staining in the lungs, all of which can contribute to barrier dysfunction [9]. Considering this, we sought to test the hypothesis that cell death, specifically apoptosis, is a major contributor to CFH-mediated lung microvascular endothelial barrier dysfunction.

Materials and Methods

Cell Culture

Primary human lung microvascular endothelial cells (HLMVECs) from several donors were purchased from PromoCell (C-12281) and grown in Endothelial Cell Basal Media MV2 (Promocell C-22221) supplemented with Growth Medium MV2 Supplement Pack (Promocell C-39221) and penicillin-streptomycin. Cells were seeded on plates coated with 0.1% gelatin and incubated at 37°C in a 5% CO2 humidified incubator to 2–3 days past confluence before being used for experimental assays.

Transendothelial Electrical Resistance (TER)

HLMVECs were seeded onto 8W10E+ PET electrode plates. The transendothelial electrical resistance (TER) was measured using an Electric Cell-Substrate Impedance Sensing (ECIS) system (Applied Biophysics, Inc). TER was used as an indicator of barrier function and was continuously measured over time. Cells were treated with vehicle control or native human cell-free hemoglobin (CFH; Cell Sciences Hb3+, CSI19668B) that was dissolved in PBS and sterile filtered, with or without a one-hour pretreatment of different cell death inhibitors, including an apoptosis inhibitor (Z-VAD-FMK, Sigma V116; 0.1 μM), necrosis inhibitor (IM-54, Millipore 480060; 5 μM), necroptosis inhibitor (necrostatin-1, Sigma N9037; 5 μM), ferroptosis inhibitor (ferrostatin-1, Sigma SML0583; 1 μM), and autophagy inhibitor (3-methyladenine, Sigma M9281; 0.2 mM). The resistance at a given time was subtracted from the baseline resistance (T=0) of each well and averaged to determine change in TER (TER Δ).

XPerT Assay

The XPerT endothelial permeability assay was adapted from previously published methods [10]. Biotinylated gelatin was prepared by mixing a 5.7 mg/mL solution of EZ-Link-NHS-LC-LC-Biotin (Thermo 21338) in DMSO with an 11.1 mg/mLsolution of gelatin in 0.1 M bicarbonate buffer at a ratio of 1:10. The resulting solution was allowed to sit at room temperature for one hour to equilibrate then further diluted with bicarbonate buffer to make a final solution of 0.5% gelatin. The biotinylated gelatin was added to a sterile 96 well, PS, F-bottom, μClear® black CellStar® TC microplate (Greiner Bio-One 655090) and incubated overnight at 4°C before seeding with HLMVECs. Two days post-confluence, cells were treated with CFH (0.5 mg/mL) or vehicle control. In order to visualize gap formation between cells, exposed biotinylated gelatin was labeled with Streptavidin-Alexa Fluor 647 (Invitrogen S21374) at 2.5 mg/mL in complete media for three minutes at room temperature, washed once with media, fixed with 4% paraformaldehyde at room temperature for ten minutes, rinsed twice with PBS, and stained with Hoechst (100 ng/mL, 30–40 minutes). Images were captured using Lionheart FX (BioTek Instruments, Inc) at 4X magnification in DAPI (377,447) and CY5 (628, 685). Images were analyzed using Gen5 Image+ software version 3.10. Montage images were stitched using Linear Blend of DAPI channel. Cellular analysis to count total number of cells included those between 5–50 μm meeting fluorescent threshold of 5,000 (DAPI). Image statistics were used to measure the total fluorescent area (CY5>10,000).

Draq7 Assay

HLMVECs were seeded on a sterile 96 well microplate (Greiner Bio-One 655090) and treated with CFH or vehicle control. Cells were exposed to the Draq7 (Abcam ab109202) stain at 1.5 μM diluted in complete media for 10 minutes, then Hoechst (100 ng/mL, 35 minutes) Live cells were imaged using a Lionheart FX (BioTek) at 4X magnification in DAPI (377,447) and CY5 (628, 685) channels. Images were analyzed using Gen5 Image+ software version 3.10. Montage images were stitched using Linear Blend of DAPI channel. Image preprocessing was applied using Auto Background Flattening of DAPI channel. Cellular analysis to count total number of cells included those between 5–50 μm meeting fluorescent threshold of 5,000 (DAPI); cells meeting fluorescence threshold (CY5>10,000) were counted as Draq7+.

TUNEL Assay

Fluorescein In Situ Cell Death Detection Kit (Roche Diagnostics 11684795910) was used for the terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay. HLMVECs were seeded onto a sterile 96 well microplate (Greiner Bio-One 655090) and treated with CFH or vehicle control. Cells were fixed with 4% paraformaldehyde for ten minutes at room temperature, rinsed twice with PBS, stained with TUNEL reaction mixture for at least one hour at 37°C, rinsed three times with PBS, labeled with Hoechst (100 ng/mL, 35 minutes), and left in PBS for imaging. Images were captured using Lionheart FX (BioTek) at 20X magnification in DAPI (377,447) and GFP (469, 525) channels. Images were analyzed using Gen5 Image+ software version 3.10. Image preprocessing was applied using Auto Background Flattening of DAPI channel. Cellular analysis to count total number of cells included those between 5–30 μm meeting fluorescent threshold of 5,000 (DAPI); cells meeting fluorescence threshold (GFP>5,000) were counted as TUNEL+.

Western Blot

Whole cell lysate collected from HLMVECs (CFH vs control) using RIPA buffer supplemented with protease (cOmplete™ Protease Inhibitor Cocktail, Roche) and phosphatase (PhosSTOP™, Roche) inhibitors were mixed with Bolt™ Sample Reducing Agent (Novex B0009) and Bolt™ LDS Sample Buffer (Novex B0007), heated for 10 minutes at 70°C, and loaded into a 4–12% Bolt™ Bis-Tris gel (Thermo NW04120BOX) at 400 V for 30 minutes. The gel was rinsed in TBS and proteins were transferred to PVDF membrane (iBlot Transfer Stack, Thermo IB24002) using iBlot 2 Gel Transfer Device (Method P0). The membrane was blocked in TBS Odyssey Blocking Buffer (Li-Cor 927–60001) for one hour and exposed to rabbit monoclonal cleaved caspase-3 (CST #9664S; 1:1000) and mouse monoclonal β-actin (CST #3700S; 1:2000) in TBS OBB (0.15% Tween-20) overnight (4°C). The next day, membrane was washed with 0.1% Tween-20 in TBS, incubated in IRDye® 800CW Goat anti-Rabbit (Li-Cor 926–3221, 1:20000) and IRDye® 680LT Goat anti-Mouse (Li-Cor 926–68020, 1:20000; 0.15% Tween-20; 0.02% SDS; TBS OBB), washed again, and imaged on Odyssey Li-Cor CLx. Densitometry analysis was performed using Image Studio Lite Version 5.2. Band density of cleaved caspase-3 was normalized to β-actin and average normalized band density from three independent experiments is displayed. Treatment of HLMVECs with staurosporine (1 μM, 4 hours; Abcam ab120056) was used as a positive apoptosis control for detection of cleaved caspase-3 via western blot.

Statistical Analysis

Statistical analysis was performed using GraphPad Prism (9.0.0) with significance set at α = 0.05. Data was analyzed using unpaired t test to compare two groups or one-way ANOVA with Tukey’s multiple comparisons post hoc analysis to compare more than two groups. Data is represented by box and whisker plots (box represents 25th to 75th percentile, line represents median, and whiskers extend from min to max).

Results:

CFH induces human lung microvascular endothelial barrier dysfunction

To test the effect of CFH on human lung microvascular endothelial barrier function, HLMVECs were treated with 0.5 mg/ml CFH. Electric cell-substrate impedance sensing (ECIS) was used to measure electrical resistance across the cell monolayer, indicating barrier tightness. HLMVECs stimulated with CFH displayed a significant decrease in resistance compared to control after six hours (Fig. 1A and B). XPerT assay, which utilizes fluorescent streptavidin binding to exposed biotinylated gelatin to reveal gaps in a cell monolayer, was used to visualize intercellular gap formation of HLMVECs exposed to CFH. Treatment of HLMVECs with CFH demonstrated a significant increase in fluorescent area at six hours (Fig. 1C and D). Pretreatment with haptoglobin, the high-affinity hemoglobin scavenger, prevented the drop in TER of HLMVECs stimulated with CFH (Fig. 1E).

Figure 1. CFH induces human lung microvascular endothelial barrier dysfunction.

Figure 1.

Open in a new tab

(A) Representative TER tracing and (B) quantification of average TER drop shows significant decrease in HLMVEC barrier dysfunction (n=9, p<0.0001) after six hours of CFH (0.5 mg/mL) stimulation. (C) Representative images (blue=nuclei, red=AF647-streptavidin) and (D) quantification of AF647-streptavidin fluorescent area show a significant increase in HLMVEC cell-cell gap formation utilizing XPerT assay (n=13, p=0.0007). (E) Average TER drop of HLMVECs stimulated with CFH (0.5 mg/mL) with or without hemoglobin scavenger haptoglobin (0.5 mg/mL) shows haptoglobin prevents HLMVEC barrier dysfunction after 24 hours (n=6–9, p<0.0001).

CFH decreases cell viability

To determine if CFH causes cell death in vitro, HLMVECs were stimulated with 0.5 mg/ml CFH for six hours and stained with Draq7, a membrane impermeable nuclear stain. There were no differences in total cell count, but a small but significant increase in both number and percentage of Draq7-positive cells was observed in CFH-treated cells (3.4%) compared to control (1.6%), indicating cell membrane compromise (Fig. 2A and B).

Figure 2. CFH decreases cell viability.

Figure 2.

Open in a new tab

(A) Representative images (blue=nuclei, red=Draq7+; arrows point to Draq7+-cells) and quantification of (B) total cell count (ns), (C) Draq7-positive cell count (p=0.01), and (D) percentage of Draq7-positive cells (p=0.03) show a significant increase in cell membrane compromise of HLMVECs stimulated with CFH (0.5 mg/mL) for 6 hours (n=21–22). ns=not significant

CFH-mediated endothelial barrier dysfunction is not caused by apoptosis

We hypothesized that the decrease in cell viability was caused by apoptosis. However, there was no significant difference in TUNEL staining (Fig. 3A–C) or cleaved caspase-3 (Fig. 3D and E) after HLMVECs were exposed to CFH for six hours. Moreover, pretreatment of HLMVEC monolayers with apoptosis inhibitor Z-VAD-FMK (0.1 μm, one hour) did not block CFH-induced endothelial barrier dysfunction measured by ECIS (Fig. 3F and G).

Figure 3. CFH-mediated endothelial barrier dysfunction is not caused by apoptosis.

Figure 3.

Open in a new tab

Quantification of (A) total cell count, (B) TUNEL-positive cell count, and (C) percentage of TUNEL-positive cells show no significant increase in apoptosis of HLMVECs stimulated with CFH (0.05 mg/mL) after six hours (n=9). (D,E) Western blot shows no increase in cleaved caspase-3 of HLMVECs stimulated with CFH (0.05 mg/mL) after six hours (n=3). (F) Representative TER tracing and (G) quantification of average TER drop shows no prevention of HLMVEC barrier dysfunction with apoptosis inhibitor Z-VAD-FMK (ZVAD, 0.1 μM, 1-hour pre-treatment; n=8–12). *=p<0.05 vs. Ctrl. ns=not significant

CFH-mediated endothelial barrier dysfunction is not caused by necrosis, necroptosis, ferroptosis, or autophagy

To determine whether other cell death pathways contributed to the cytotoxicity observed with Draq7 staining, several other inhibitors of pathways mediating cell death were tested for ability to prevent barrier dysfunction. However, necrosis inhibitor IM-54 (5 μM; Fig. 4A and B), necroptosis inhibitor necrostatin-1 (5 μM; Fig. 4C and D), ferroptosis inhibitor ferrostatin-1 (1 μM; Fig. 4E and F), and autophagy inhibitor 3-methyladenine (0.2 mM; Fig. 4G and H) were unable to prevent the barrier dysfunction induced by CFH.

Figure 4. CFH-mediated endothelial barrier dysfunction is not caused by necrosis, necroptosis, ferroptosis, or autophagy.

Figure 4.

Open in a new tab

(A,C,E,G) Representative TER tracing and (B,D,F,H) quantification of average TER drop (n=8–13) shows no prevention of HLMVEC barrier dysfunction with one-hour pre-treatment of (A,B) necrosis inhibitor IM-54 (5 μM), (C,D) necroptosis inhibitor necrostatin-1 (NS1, 5 μM), (E,F) ferroptosis inhibitor ferrostatin-1 (FS1, 1 μM), or (G,H) autophagy inhibitor 3-methyladenine (3MA, 0.2 mM). *=p<0.05 vs. Ctrl. ns=not significant

Discussion

In this study, we show that CFH causes human lung microvascular endothelial barrier dysfunction that can be blocked by the hemoglobin scavenger, haptoglobin. Though CFH caused a small but significant increase in cytotoxicity, CFH-mediated lung microvascular endothelial barrier dysfunction was not caused by known cell death pathways including apoptosis, necrosis, necroptosis, ferroptosis, and autophagy.

Our group and others have shown that higher circulating levels of CFH contribute to organ injury and poor outcomes in several hemolytic and inflammatory pathologies [1, 11–13]. Moreover, increased circulating levels of CFH in animal models such as sepsis [9], transfusion-induced acute lung injury [14], pulmonary hypertension [12], and sickle cell disease [15] contribute to lung endothelial barrier dysfunction and microvascular hyperpermeability. We previously showed direct evidence of CFH-mediated vascular permeability using an ex vivo perfused human lung model; hemoglobin infusion resulted in increased lung weight and Evans blue-labeled albumin leak [13]. Several studies have examined the endothelial barrier disrupting effects of CFH in animal models of hemoglobin infusion [9, 16, 17] and via direct hemoglobin stimulation in vitro [8, 18–22]. Though many of these studies offer clues to the effects of CFH on mediating endothelial injury, a complete understanding of the mechanisms involved in CFH-mediated microvascular endothelial barrier dysfunction remains to be elucidated.

Our initial investigation into cell death pathways was in an effort to explain the significant increase previously observed in TUNEL-positive staining of lung tissue in a mouse model of sepsis with increased circulating CFH and in an immortalized line of human lung microvascular endothelial cells (HULEC-5a) [9]. TUNEL specifically measures DNA fragmentation, a main characteristic of apoptotic cell death [10]. In this study, we did not observe a significant increase in TUNEL staining of HLMVECs in response to CFH treatment in vitro. To further rule out apoptosis, we measured cleaved caspase-3, an accepted apoptotic marker involved in several apoptotic processes [23], via western blot and saw no difference in cleaved caspase-3 after CFH stimulation. The discrepancies in observed apoptosis between this study and our previous work could be explained by the differences in characteristics between the immortalized cell line and primary cells, or by the complexity of the in vivo sepsis model previously used, in which there are many additional factors such as circulating cytokines and immune cells, and there is potential for hemoglobin to be oxidized to its more injurious ferryl (4+) form.

In addition to the cell death pathways studied herein, there are several other known forms of cell death—such as pyroptosis, autosis and more—and likely others that have not yet been discovered [24–26]. It is possible that one of these pathways, or a completely novel mechanism, is triggered in response to CFH leading to increased barrier permeability. Notably, however, CFH-induced increases in endothelial barrier dysfunction were observed at approximately two hours, but no difference in cell viability was observed until after six hours. This indicates that even though a small amount of endothelial cell death may be induced by CFH over time, it is not the primary driver of endothelial barrier dysfunction. Future studies into precise signaling mechanisms in the endothelium may offer clues to determine which known or potentially novel cell death pathways might be activated in response to CFH.

It has also been reported that the plasma membrane can be disrupted independent of cell death with repair mechanisms protecting against cell lysis [27, 28]. Potential mechanisms involved in plasma membrane repair include patching or resealing via protein aggregation, endocytosis or shedding of compromised membrane, and exocytosis or release of vesicles; many of these mechanisms involve calcium influx and cytoskeletal rearrangement [27, 29–32]. Since the Draq7 dye used in this study is a non-specific cell impermeable nuclear stain, it is possible that the increase in Draq7-positive cells observed reflects not cell death, but cell membrane compromise. Oxidized forms of hemoglobin have been shown to induce cytoskeletal rearrangement and upregulation of adhesion molecules in endothelial cells [33, 34]; these pathways can decrease barrier function while also disrupting the plasma membrane and are possible mechanisms for hemoglobin-induced injury in lung microvascular endothelial cells.

Despite initial evidence from murine models, we have determined that direct CFH-induced human lung microvascular endothelial barrier dysfunction is not caused by a common cell death mechanism. The precise signaling mechanisms leading to CFH-mediated endothelial barrier dysfunction remain to be elucidated and will be the focus of our future studies.

Highlights.

  • Cell-free hemoglobin (CFH) causes human lung microvascular EC barrier dysfunction

  • Haptoglobin rescues CFH-mediated HLMVEC barrier dysfunction

  • CFH induces a modest but significant decrease in cell viability after six hours

  • CFH-mediated HLMVEC barrier dysfunction is not caused by common forms of cell death

Acknowledgments

Funding

This work was supported by National Institutes of Health HL135849 to LBW and JAB; HL103836 to LBW; HL094296 to JEM.

Abbreviations:

CFH

cell-free hemoglobin

ARDS

acute respiratory distress syndrome

ROS

reactive oxygen species

HLMVECs

human lung microvascular endothelial cells

TER

transendothelial electrical resistance

ECIS

Electric Cell-Substrate Impedance Sensing

TUNEL

terminal deoxynucleotidyl transferase dUTP nick end labeling

Footnotes

Declaration of competing interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Please note that all Biochemical and Biophysical Research Communications authors are required to report the following potential conflicts of interest with each submission. If applicable to your manuscript, please provide the necessary declaration in the box above.

(1) All third-party financial support for the work in the submitted manuscript.

(2) All financial relationships with any entities that could be viewed as relevant to the general area of the submitted manuscript.

(3) All sources of revenue with relevance to the submitted work who made payments to you, or to your institution on your behalf, in the 36 months prior to submission.

(4) Any other interactions with the sponsor of outside of the submitted work should also be reported. (5) Any relevant patents or copyrights (planned, pending, or issued).

(6) Any other relationships or affiliations that may be perceived by readers to have influenced, or give the appearance of potentially influencing, what you wrote in the submitted work. As a general guideline, it is usually better to disclose a relationship than not.

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  • [1].Janz DR, Ware LB, The role of red blood cells and cell-free hemoglobin in the pathogenesis of ARDS, Journal of intensive care, 3 (2015) 20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [2].Kerchberger VE, Bastarache JA, Shaver CM, Nagata H, McNeil JB, Landstreet SR, Putz ND, Yu W-K, Jesse J, Wickersham NE, Sidorova TN, Janz DR, Parikh CR, Siew ED, Ware LB, Haptoglobin-2 variant increases susceptibility to acute respiratory distress syndrome during sepsis, JCI Insight, 4 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [3].Huppert LA, Matthay MA, Ware LB, Pathogenesis of Acute Respiratory Distress Syndrome, Seminars in respiratory and critical care medicine, 40 (2019) 31–39. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [4].Effenberger-Neidnicht K, Hartmann M, Mechanisms of Hemolysis During Sepsis, Inflammation, 41 (2018) 1569–1581. [DOI] [PubMed] [Google Scholar]
  • [5].Bozza MT, Jeney V, Pro-inflammatory Actions of Heme and Other Hemoglobin-Derived DAMPs, Frontiers in Immunology, 11 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [6].Adamzik M, Hamburger T, Petrat F, Peters J, de Groot H, Hartmann M, Free hemoglobin concentration in severe sepsis: methods of measurement and prediction of outcome, Critical Care, (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [7].Shaver CM, Wickersham N, McNeil JB, Nagata H, Sills G, Kuck JL, Janz DR, Bastarache JA, Ware LB, Cell-Free Hemoglobin-mediated Increases in Vascular Permeability. A Novel Mechanism of Primary Graft Dysfunction and a New Therapeutic Target, Annals of the American Thoracic Society, 14 (2017) S251–S252. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [8].Kuck JL, Bastarache JA, Shaver CM, Fessel JP, Dikalov SI, May JM, Ware LB, Ascorbic acid attenuates endothelial permeability triggered by cell-free hemoglobin, Biochemical and Biophysical Research Communications, 495 (2018) 433–437. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [9].Meegan JE, Shaver CM, Putz ND, Jesse JJ, Landstreet SR, Lee HNR, Sidorova TN, McNeil JB, Wynn JL, Cheung-Flynn J, Komalavilas P, Brophy CM, Ware LB, Bastarache JA, Cell-free hemoglobin increases inflammation, lung apoptosis, and microvascular permeability in murine polymicrobial sepsis, PLOS ONE, 15 (2020) e0228727. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [10].Dubrovskyi O, Birukova AA, Birukov KG, Measurement of local permeability at subcellular level in cell models of agonist-and ventilator-induced lung injury, Laboratory Investigation, (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [11].Janz DR, Bastarache JA, Peterson JF, Sills G, Wickersham N, May AK, Roberts LJ, Ware LB, Association between cell-free hemoglobin, acetaminophen, and mortality in patients with sepsis: An observational study, Critical Care Medicine, (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [12].Rafikova O, Williams ER, McBride ML, Zemskova M, Srivastava A, Nair V, Desai AA, Langlais PR, Zemskov E, Simon M, Mandarino LJ, Rafikov R, Hemolysis-induced Lung Vascular Leakage Contributes to the Development of Pulmonary Hypertension, Am J Respir Cell Mol Biol, 59 (2018) 334–345. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [13].Shaver CM, Wickersham N, McNeil JB, Nagata H, Miller A, Landstreet SR, Kuck JL, Diamond JM, Lederer DJ, Kawut SM, Palmer SM, Wille KM, Weinacker A, Lama VN, Crespo MM, Orens JB, Shah PD, Hage CA, Cantu E, Porteous MK, Dhillon G, McDyer J, Bastarache JA, Christie JD, Ware LB, Cell-free hemoglobin promotes primary graft dysfunction through oxidative lung endothelial injury, JCI Insight, 3 (2018) 1–12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [14].Stapley R, Rodriguez C, Oh JY, Honavar J, Brandon A, Wagener BM, Marques MB, Weinberg JA, Kerby JD, Pittet JF, Patel RP, Red blood cell washing, nitrite therapy, and antiheme therapies prevent stored red blood cell toxicity after trauma-hemorrhage, Free Radical Biology and Medicine, 85 (2015) 207–218. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [15].Ghosh S, Tan F, Ofori-Acquah SF, Spatiotemporal dysfunction of the vascular permeability barrier in transgenic mice with sickle cell disease, Anemia, 2012 (2012) 582018–582018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [16].Butt OI, Buehler PW, D’Agnillo F, Blood-brain barrier disruption and oxidative stress in guinea pig after systemic exposure to modified cell-free hemoglobin, The American journal of pathology, 178 (2011) 1316–1328. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [17].Fu Z, Chen Y, Qin F, Yang S, Deng X, Ding R, Feng L, Li W, Zhu J, Increased activity of Rho kinase contributes to hemoglobin-induced early disruption of the blood-brain barrier in vivo after the occurrence of intracerebral hemorrhage, International Journal of Clinical and Experimental Pathology, (2014). [PMC free article] [PubMed] [Google Scholar]
  • [18].Dull RO, DeWitt BJ, Dinavahi R, Schwartz L, Hubert C, Pace N, Fronticelli C, Quantitative assessment of hemoglobin-induced endothelial barrier dysfunction, Journal of applied physiology (Bethesda, Md. : 1985), 97 (2004) 1930–1937. [DOI] [PubMed] [Google Scholar]
  • [19].Jana S, Meng F, Hirsch RE, Friedman JM, Alayash AI, Oxidized Mutant Human Hemoglobins S and E Induce Oxidative Stress and Bioenergetic Dysfunction in Human Pulmonary Endothelial Cells, Frontiers in Physiology, 8 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [20].Kim J, Nguyen TTT, Li Y, Zhang CO, Cha B, Ke Y, Mazzeffi MA, Tanaka KA, Birukova AA, Birukov KG, Contrasting effects of stored allogeneic red blood cells and their supernatants on permeability and inflammatory responses in human pulmonary endothelial cells, Am J Physiol Lung Cell Mol Physiol, 318 (2020) L533–l548. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [21].Lisk C, Kominsky D, Ehrentraut S, Bonaventura J, Nuss R, Hassell K, Nozik-Grayck E, Irwin DC, Hemoglobin-induced endothelial cell permeability is controlled, in part, via a myeloid differentiation primary response gene-88-dependent signaling mechanism, American Journal of Respiratory Cell and Molecular Biology, 49 (2013) 619–626. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [22].Potor L, Nagy P, Mehes G, Hendrik Z, Jeney V, Petho D, Vasas A, Palinkas Z, Balogh E, Gyetvai A, Whiteman M, Torregrossa R, Wood ME, Olvaszto S, Nagy P, Balla G, Balla J, Hydrogen Sulfide Abrogates Hemoglobin-Lipid Interaction in Atherosclerotic Lesion, Oxidative medicine and cellular longevity, 2018 (2018) 3812568. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [23].Porter AG, Jänicke RU, Emerging roles of caspase-3 in apoptosis, Cell death and differentiation, 6 (1999) 99–104. [DOI] [PubMed] [Google Scholar]
  • [24].Pober JS, Min W, Bradley JR, Mechanisms of endothelial dysfunction, injury, and death, Annual review of pathology, 4 (2009) 71–95. [DOI] [PubMed] [Google Scholar]
  • [25].Galluzzi L, Vitale I, Aaronson SA, Abrams JM, Adam D, Agostinis P, Alnemri ES, Altucci L, Amelio I, Andrews DW, Annicchiarico-Petruzzelli M, Antonov AV, Arama E, Baehrecke EH, Barlev NA, Bazan NG, Bernassola F, Bertrand MJM, Bianchi K, Blagosklonny MV, Blomgren K, Borner C, Boya P, Brenner C, Campanella M, Candi E, Carmona-Gutierrez D, Cecconi F, Chan FKM, Chandel NS, Cheng EH, Chipuk JE, Cidlowski JA, Ciechanover A, Cohen GM, Conrad M, Cubillos-Ruiz JR, Czabotar PE, D’Angiolella V, Dawson TM, Dawson VL, De Laurenzi V, De Maria R, Debatin K-M, DeBerardinis RJ, Deshmukh M, Di Daniele N, Di Virgilio F, Dixit VM, Dixon SJ, Duckett CS, Dynlacht BD, El-Deiry WS, Elrod JW, Fimia GM, Fulda S, García-Sáez AJ, Garg AD, Garrido C, Gavathiotis E, Golstein P, Gottlieb E, Green DR, Greene LA, Gronemeyer H, Gross A, Hajnoczky G, Hardwick JM, Harris IS, Hengartner MO, Hetz C, Ichijo H, Jäättelä M, Joseph B, Jost PJ, Juin PP, Kaiser WJ, Karin M, Kaufmann T, Kepp O, Kimchi A, Kitsis RN, Klionsky DJ, Knight RA, Kumar S, Lee SW, Lemasters JJ, Levine B, Linkermann A, Lipton SA, Lockshin RA, López-Otín C, Lowe SW, Luedde T, Lugli E, MacFarlane M, Madeo F, Malewicz M, Malorni W, Manic G, Marine J-C, Martin SJ, Martinou J-C, Medema JP, Mehlen P, Meier P, Melino S, Miao EA, Molkentin JD, Moll UM, Muñoz-Pinedo C, Nagata S, Nuñez G, Oberst A, Oren M, Overholtzer M, Pagano M, Panaretakis T, Pasparakis M, Penninger JM, Pereira DM, Pervaiz S, Peter ME, Piacentini M, Pinton P, Prehn JHM, Puthalakath H, Rabinovich GA, Rehm M, Rizzuto R, Rodrigues CMP, Rubinsztein DC, Rudel T, Ryan KM, Sayan E, Scorrano L, Shao F, Shi Y, Silke J, Simon H-U, Sistigu A, Stockwell BR, Strasser A, Szabadkai G, Tait SWG, Tang D, Tavernarakis N, Thorburn A, Tsujimoto Y, Turk B, Vanden Berghe T, Vandenabeele P, Vander Heiden MG, Villunger A, Virgin HW, Vousden KH, Vucic D, Wagner EF, Walczak H, Wallach D, Wang Y, Wells JA, Wood W, Yuan J, Zakeri Z, Zhivotovsky B, Zitvogel L, Melino G, Kroemer G, Molecular mechanisms of cell death: recommendations of the Nomenclature Committee on Cell Death 2018, Cell Death & Differentiation, 25 (2018) 486–541. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [26].Chen Y, Hua Y, Li X, Arslan IM, Zhang W, Meng G, Distinct Types of Cell Death and the Implication in Diabetic Cardiomyopathy, Front Pharmacol, 11 (2020) 42–42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [27].Andrews NW, Corrotte M, Plasma membrane repair, Current Biology, 28 (2018) R392–R397. [DOI] [PubMed] [Google Scholar]
  • [28].Dias C, Nylandsted J, Plasma membrane integrity in health and disease: significance and therapeutic potential, Cell discovery, 7 (2021) 4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [29].Jimenez AJ, Perez F, Plasma membrane repair: the adaptable cell life-insurance, Current Opinion in Cell Biology, 47 (2017) 99–107. [DOI] [PubMed] [Google Scholar]
  • [30].Miyake K, McNeil PL, Suzuki K, Tsunoda R, Sugai N, An actin barrier to resealing, Journal of cell science, 114 (2001) 3487–3494. [DOI] [PubMed] [Google Scholar]
  • [31].Andrews AM, Lutton EM, Merkel SF, Razmpour R, Ramirez SH, Mechanical Injury Induces Brain Endothelial-Derived Microvesicle Release: Implications for Cerebral Vascular Injury during Traumatic Brain Injury, Frontiers in cellular neuroscience, 10 (2016) 43. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [32].Miyake K, McNeil PL, Mechanical injury and repair of cells, Crit Care Med, 31 (2003) S496–501. [DOI] [PubMed] [Google Scholar]
  • [33].Posta N, Csosz E, Oros M, Petho D, Potor L, Kallo G, Hendrik Z, Sikura KE, Mehes G, Toth C, Posta J, Balla G, Balla J, Hemoglobin oxidation generates globin-derived peptides in atherosclerotic lesions and intraventricular hemorrhage of the brain, provoking endothelial dysfunction, Laboratory investigation; a journal of technical methods and pathology, (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • [34].Silva G, Jeney V, Chora A, Larsen R, Balla J, Soares MP, Oxidized hemoglobin is an endogenous proinflammatory agonist that targets vascular endothelial cells, The Journal of biological chemistry, 284 (2009) 29582–29595. [DOI] [PMC free article] [PubMed] [Google Scholar]

RESOURCES