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. Author manuscript; available in PMC: 2015 Jun 1.
Published in final edited form as: J Thorac Cardiovasc Surg. 2013 Nov 13;147(6):1939–1945. doi: 10.1016/j.jtcvs.2013.09.054

Aprotinin, but not epsilon aminocaproic acid and tranexamic acid, exerts neuroprotection against excitotoxic injury in an in vitro neuronal cell culture model

Zhaohui Lu 1,3, Ludmila Korotcova 1,2, Akira Murata 1,2, Nobuyuki Ishibashi 1,2,4, Richard A Jonas 1,2,4
PMCID: PMC4019727  NIHMSID: NIHMS541298  PMID: 24237885

Abstract

Objective

Lack of availability of aprotinin has resulted in increased clinical use of the alternative antifibrinolytic agents epsilon aminocaproic acid (EACA) and tranexamic acid (TXA) which are known to be associated with an increased risk of seizures. In contrast aprotinin has previously been demonstrated to be neuroprotective through suppression of excitotoxicity-mediated neuronal degeneration via the extracellular plasminogen/plasmin system. We compared the impact of antifibrinolytic agents on neuronal and mixed glial/neuronal cell cultures.

Methods

Mixed cortical cultures containing neuronal and glial cells were prepared from fetal mice and plated on a layer of confluent astrocytes from postnatal pups. Primary neuronal culture was obtained from the same gestational stage and plated in multiwall vessels. Slowly triggered excitotoxicity was induced by 24-hour exposure to 12.5 mM N-methyl-D-aspartate (NMDA). Apoptotic neuronal cell death was induced by exposure of primary neural cultures to 24 hours of serum deprivation.

Results

Compared to NMDA alone, no significant changes in cell death were observed for any dose of TXA or EACA in mixed cultures. Conversely, a clinical dose of aprotinin significantly reduced cell death by -31% on average. Aprotinin reduced apoptotic neuronal cell death from 75% to 37.3%, and 34.1% at concentrations of 100 and 200 KIU/mL, and significantly decreased neuronal nuclear damage. These concentrations of aprotinin significantly inhibited caspase 9 and 3/7 activations. 250 KIU/ml aprotinin exerted maximal protection on primary cortical neurons.

Conclusions

In contrast to aprotinin, EACA and TXA exert no protective effect against excitotoxic neuronal injury that can occur during cardiac surgery.

Low flow cardiopulmonary bypass and deep hypothermic circulatory arrest are used to facilitate pediatric cardiac surgery although they carry a risk of ischemic brain damage. One mechanism of ischemic neuronal cell death is “glutamatergic excitotoxcity”.1 Initially an ischemic insult impairs glutamate transport at the postsynaptic level resulting in high extracellular glutamate levels1, 2 resulting in excessive calcium influx eventually leading to neuronal cell death1.

Studies of excitotoxicity-mediated neuronal degeneration have identified the extracellular plasminogen/plasmin system as an important aggravating element, eg through activation of intrinsic tissue-plasminogen activator.3, 4 Aprotinin, which is used as an anti-fibrinolytic agent during cardiac surgery inhibits plasminogen activation. We previously demonstrated that aprotinin has neuroprotective effects against glutamatergic excitotoxicity.5 It also protects endothelial function in the brain.6 Lysine analogues such as epsilon aminocaproic acid (EACA) and tranexamic acid (TXA) block the binding of plasminogen to fibrin and plasminogen activation and transformation to plasmin.7 The lysine analogues, therefore, also have the potential for neuroprotection through their inhibition of plasminogen activation.

In the present study, we first used a neuronal cell culture model to examine whether antifibrinolytic lysine analogues also are neuroprotective against glutamatergic excitotoxicity. We then assessed the anti-apoptotic effects of aprotinin. Finally, we studied the mechanisms underlying the neuroprotective effects of aprotinin and investigated the optimal dose for clinical application during cardiac surgery.

Methods

Primary cortical neuron cultures

Primary cortical neuron cultures containing less than 5% astrocytes were obtained from fetal mice at 13– 15d gestation (Charles River, Germantown, MD). Dissociated cortical cells in plating medium of media stock (MS: Dulbecco's modified Eagle's medium with 25 mM glucose; SIGMA, St. Louis, MO) supplemented with 5% fetal bovine serum (GIBCO, Carlsbad, CA), 5% horse serum (GIBCO, Carlsbad, CA) and 2 mM glutamine (SIGMA, St. Louis, MI) were plated in 24-well plates coated with poly-D-lysine (0.1 mg/dl; Invitrogen, Carlsbad, CA) and laminin (0.02 mg/ml; Invitrogen, Carlsbad, CA). After 3 days in vitro (DIV), non-neuronal cell division was halted by exposure to 10 μM cytosine arabinoside (Ara-C; SIGMA, St. Louis, MO). There was no further exchange of the media except adding DMEM for evaporation. All cultures were kept at 37°C in a humidified 5% CO2 incubator. Cultures were used after 7 DIV for serum deprivation (SD). We performed all experiments in compliance with the NIH Guide for the Care and Use of Laboratory animals. The study was approved by the Institutional Animal Care and Use Committee of the Children's National Medical Center.

Glial cell cultures

Glial cell cultures were prepared from 1 to 3-day-old postnatal mice (Swiss Webster mice; Charles River, Germantown, MD). Dissociated cortical cells were plated in 24-well plates previously coated with poly-D-lysine using a plating medium of MS supplemented with 10% horse serum, 10% fetal bovine serum (Gibco, Carlsbad, CA), and 2mmol/L glutamine. Cultures were kept at 37°C in a humidified 5% CO2-containing atmosphere until they reached confluence 7 to 14 days in vitro. Confluent cultures were then used as a support for mixed cultures.

Mixed cortical cultures

Mixed cortical cultures containing both neurons and astrocytes were prepared from fetal mice at 14.5 days gestation. Dissociated cortical cells were plated in 24 wells on a layer of confluent astrocytes, using MS supplemented with 5% horse serum, 5% fetal bovine serum, and 2mmol/L glutamine. After 7 days in vitro, non-neuronal cell division was halted by 3 days of exposure to 10 mM cytosine arabinoside (Ara-C; Sigma). Subsequent partial medium replacement was performed twice per week, and after 12 days in vitro, cultures were shifted to a maintenance medium identical to the plating medium but lacking serum, because neurons survive without it. Experiments were performed on cortical cultures after 13 to 14 days in vitro.

Excitotoxicity

Slowly triggered excitotoxicity was induced at 37°C by 24-hour exposure to 12.5μM N-Methyl-D-aspartic acid (NMDA) as an excitotoxin in medium stock supplemented with 10mM glycine. TXA, EACA, and aprotinin was co-applied at 3 different concentrations (Low dose, Clinical dose, High dose) with the excitotoxin and left for 24hrs in the bathing medium. The clinical therapeutic concentration of each agent is 0.8mM for TXA, 1mM for EACA8, and 250KIU/ml for aprotinin9. Low and high dose were 1/3 and 3 times the clinical dose, respectively. Since it has been recommended to maintain the plasma TXA concentration above 0.8mM for high-risk bleeding patients, 10 we choose 0.8mM as the clinical therapeutic concentration of TXA in this study, which corresponds with the recommended dosing of TXA 100 mg/kg as a single bolus before surgery or a regimen with a loading dose of 30 mg/kg plus additional 2 mg/kg added to the pump prime followed by 16 mg/kg/h infusion in adult patients.10

Neuronal death resulting from slowly triggered excitotoxicity was confirmed by examining cultures under phase-contrast microscopy and quantified by measurement of lactate dehydrogenase (LDH) release from damaged cells into the bathing medium 1 day after the onset of excitotoxin exposure. LDH was measured by an enzyme-linked immune sorbent assay kit (Promega, Madison, Wis). The LDH level corresponding to complete neuronal death was determined in sister cultures exposed to 100μM NMDA. Background LDH levels were determined in sister cultures with sham wash as a control and subtracted from experimental values to yield the signal specific for experimentally induced injury.

Serum Deprivation (SD)

Apoptosis was induced by SD for 24hr at 7 DIV in the same fashion described previously.4 In brief each cell culture was rinsed; then, aprotinin at tested concentrations was applied to the culture medim without any serum. One group was treated with normal neurobasal medium after the rinse as control (Control), and another was treated with DMEM without any serum and aprotinin (No-Aprotinin). Secondary NMDA receptor activation was blocked by the addition of MK-801 at a final concentration of 10μM to the medium. Plates were then incubated for 24hr.

Trypan Blue Staining

Trypan blue, a vital stain used to selectively color dead tissues or cells blue, was used to differentiate live neurons from dead ones. After the induction of apoptosis by SD, 100μl trypan blue prewarmed to 37°C was added to each well, and cells were incubated at 37°C for 15 min. The cells were then fixed in phosphate-buffered formalin solution (4%) for 30min at room temperature. Neurons in each dish were counted in 7 different areas using an inverted microscope. Unstained neurons with intact soma and neurites were regarded as viable, and dark-stained neurons were considered damaged. Cell counting was performed in a blinded fashion.

3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay

Viability of neurons was assessed using an MTT. Briefly, 30μl MTT was added to each well and incubated at 37°C for 3hr. The supernatant media was gently removed and cells were solubilized with 250μl dimethylsulfoxide. Then, 100μl medium was transferred to a 96-well plate and formazan reduction was detected by measuring absorbance at 570nm (OD570) with a reference wavelength of 655nm in a spectrophotometer.

Hoechst staining

The Hoechst stains belong to a family of fluorescent stains that label DNA by fluorescence. Because these fluorescent stains label DNA, they are also commonly used to visualize nuclei and mitochondria.11 Briefly, primary cortical neurons that had been treated as mentioned above were fixed in 4% paraformaldehyde for 20min, and then were stained with 20μg/ml Hoechst 33258 dye for 10min, followed by observation using a fluorescence microscope. The dye was excited at 340nm, and emission was filtered with a 510nm barrier filter. Neurons with fragmented or condensed DNA or with normal DNA were counted to quantify the apoptotic process. Data are expressed as the ratio of apoptotic neurons to total neurons.

Flow cytometry

Neuronal apoptosis was assayed by flow cytometry using an Annexin V/FITC kit, as described previously 12. After 24hr of serum deprivation, cells were washed twice with cold PBS and were detached from the bottom with trypsin-EDTA treatment for 2 to 10min at 37°C. After adding PBS with 2% FBS, neurons were harvested and centrifuged at 1500rpm for 5min. Neurons were resuspended in 1× binding buffer (10mM HEPES, 140mM NaCl, 2.5mM CaCl2) at concentration of 1 × 106 cells/ml. One hundred μl of each cell suspension was transferred to a 5ml tube and 5μl FITC-conjugated annexin V and propidium iodide (PI) (50μg/ml) was added. After 15min incubation in the dark at room temperature, 400μl binding buffer was added to each tube and cells were analyzed for annexin V binding within 1 hr with a flow cytometer.

Caspase 3/7 and 9 assay

Caspase activity was detected using Caspase 9 and 3/7 Assay Kits (Promega, Madison, WI) as per the manufacturer's protocol. Caspase 3/7 and 9 concentrations were expressed as relative light units (RLU) per mg cellular protein measured with a Reporter microplate luminometer. The background luminescence associated with the cell culture and assay reagent (blank reaction) was subtracted from the experimental values. The activities of caspases -3/7 and -9 were reported as the means of experiments conducted in triplicate.

Immunohistochemistory

To identify Caspase3-positive neural cells after 24hrs SD, cultured neurons were immunolabeled with an primary antibody to Caspase3 (Cell signaling technology, Danvers, MA) diluted in 0.1M phosphate-buffered saline (pH 7.4) containing 0.1% Triton X-100 and 5% normal goat serum for 1h at 4°C. Cells were then carefully washed in medium and incubated in the secondary antibody for 1h at 4°C, and mounted on slides. For cell counting, images were captured using a fluorescence microscope. Samples were then analyzed using ImageJ software. To determine cell density, the antibody positive cells were quantified in 5 microscopic fields from each sample (6 samples per group) in a blinded fashion.

Statistics

One-way analysis of variance (ANOVA) with Bonferroni post hoc comparisons was used to detect differences in each analysis between tested groups. Analysis of the data was performed using SPSS version 19.0 (SPSS Inc/IBM, Chicago, Ill).

Results

Aprotinin, but not Lysine Analogues, are Neuroprotective against Glutamatergic Excitotoxicity

Sham wash and the addition of two lysine analogues and aprotinin alone at high dose did not influence neuronal cells in either culture (Fig. 1a-d). Excitotoxicity induced by exposure of the cultures to 12.5μM NMDA for 24hrs resulted in acute swelling of neuronal cell bodies, followed by widespread necrotic neuronal degeneration resulting in disrupted neurons and segmentalized neurites (Fig. 1e). When EACA and TXA were co-applied in the medium with the excitotoxin, both lysine analogues did not preserve neuronal cells and neurites (Fig. 1f,g). On the other hand, the damage was significantly reduced under the condition with aprotinin (Fig. 1h). Exposure to 12.5mM NMDA caused approximately 70% neuronal cell death (Fig. 1i). There were no effects on cell death with any dose of TXA or EACA (Fig. 1i). In contrast, aprotinin reduced neuronal death in a dose-dependent fashion (Fig. 1i). Compared to NMDA alone, no significant changes in cell death were observed for clinical doses of TXA or EACA, whereas aprotinin significantly reduced cell death by -31% on average (Fig. 1j). Altogether the results indicate that, among major antifibrinolytic agents clinically used in cardiac surgery, only aprotinin has a neuroprotective effects against excitotoxic brain injury.

Fig. 1. Aprotinin, but not lysine analogues, are neuroprotective against glutamatergic excitotoxicity.

Fig. 1

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(a-d) Mixed cortical culture at 24hrs after Sham wash (a), and Sham wash with clinical dose TXA (b), EACA (c), and aprotinin (d). (e-h) Mixed cortical culture after 24hr exposure with 12.5 μM NMDA (e), and 12.5 μM NMDA with clinical dose TXA (f), EACA (g), and Aprotinin (h). (i) Percentage of cell death at 24hrs after 12.5 μM NMDA exposure with TXA, EACA, and Aprotinin. Neural cell death was identified by LDH assay measuring LDH levels in bathing medium in cultures 4. Low, clinical, and high doses of aprotinin reduced neuronal death from 70% to 56%, 39%, and 31%, respectively (F = 104.6, p < 0.0001). (j) Percentage cell death at 24hrs after 12.5 μM NMDA exposure with clinical dose of TXA, EACA, and Aprotinin compared with 12.5 μM NMDA alone. There were no significant changes in TXA (+8%, p = 0.11) and EACA (-3%, p = 0.45). A clinical dose of Aprotinin significantly reduced the death by -31% on average (p < 0.0001). NMDA; N-Methyl-D-aspartic acid, EACA; epsilon aminocaproic acid, TXA; tranexamic acid. *p < 0.001 vs. 12.5 μM NMDA by ANOVA with Bonferroni comparisons. Data are shown as mean ± SEM (n = 12).

Aprotinin Prevents Serum Deprivation-Induced Apoptosis in Cortical Neurons

In addition to the effect of aprotinin against glutamatergic excitotoxicity, our previous results suggested potential anti-apoptotic effect.13 In the present study using primary cortical neuronal culture, 100 and 200KIU/ml aprotinin significantly decreased trypan blue+ cell percentage resulting from SD-induced apoptosis, 4, 14, 15 compared with Control (Fig. 2a). Consistent with these results, cell viability using MTT assay significantly decreased at 24hrs after SD in No aprotinin; while, 100KIU/ml and 200KIU/ml aprotinin significantly increased the viability compared with No aprotinin (Fig. 2b). When Hoechst 33258 staining was performed at the same time point and damaged cells were characterized by condensed chromatin, reduced nuclear size, and nuclear fragmentation (Fig. 2c-g), it was identified that the percentage of damaged cells significantly decreased in 100 and 200KIU/ml aprotinin (25.0 ± 3.8% and 20.3 ± 3.6%, respectively) compared with No aprotinin (34.0 ± 4.7%; p < 0.01). However there were no significant differences between 50KIU/ml and No aprotinin in assessments using trypan blue, MTT, and Hoechst 33258 (Fig. 2a,b,e). The percentage of early apoptotic neurons identified by annexin V+ using flow cytometry was not significantly different between aprotinin and no aprotinin treatment after 24hrs SD. When advanced apoptosis was indicated by annexin V+ and V/PI+, 78.8% of neurons (± 10.4%) were identified as having suffered advanced apoptosis; whereas, this effect was significantly reduced by aprotinin at all three concentrations tested (50KIU/ml aprotinin, 46.1 ± 12.0%; 100KIU/ml, 37.1 ± 14.3%; 200KIU/ml, 43.5 ± 21.3%; p < 0.01). When taken together, these findings demonstrate that in addition to a neuro-protective effect against excitotoxicity, aprotinin significantly inhibits apoptosis in cortical neurons.

Fig. 2. Aprotinin prevents serum deprivation-induced apoptosis in cortical neurons.

Fig. 2

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(a) Percentage of trypan blue+ cells at 24hrs after serum deprivation. Ten percent cell death in Control supports a technical consistency in our study.35 Aprotinin at 50, 100 and 200KIU/ml decreased trypan blue+ cell percentage from 75% to 60%, 37% and 34%, respectively. (b) Cell viability using MTT assay after 24hr SD. Aprotinin at 100 and 200 KIU/ml significantly increased the viability compared with No Aprotinin (p < 0.05). There was no significant difference between No Aprotinin and 50KIU/ml Aprotinin. (c-g) Images in Hoechst 33258 staining at 24hrs after Sham wash (c) and after SD with Aprotinin at the concentration of 0 (d), 50 (e), 100 (f), and 250KIU/ml (g). *p < 0.05, **p < 0.01, ***p < 0.001 vs. Aprotinin 0KIU/ml by ANOVA with Bonferroni comparisons. Data are shown as mean ± SEM (n = 8).

Aprotinin Inhibits Apoptosis via Mitochondrial Pathway and 250KIU/ml is an Optimal Dose for Protection

To identify a pathway involving aprotinin-induced inhibition of apoptosis, caspase3/7 and 9 levels were assessed after 24hrs SD. Aprotinin significantly decreased caspase-3/7 level in a dose-dependent fashion (Fig. 3a). This dose-dependent change correlated with apoptotic neural injury identified by trypan blue and MTT assays (Fig. 2a,b), suggesting that a caspase-dependent mechanism includes an anti-apoptotic effect due to aprotinin. It was also identified that caspase-9 activity was significantly inhibited by 100 and 200KIU/ml aprotinin (Fig. 3b). When apoptosis was induced by 200nM Staurosporin, however, there were no significant differences in MTT assay between any doses of aprotinin (data not shown). It is known that staurosporin induces apoptosis through both caspase-dependent and -independent mechanisms 15; while, SD induces apoptotic cell death using a caspase-dependent mechanism via a mitochondrial pathway16. Therefore, it is likely that inhibition of apoptosis by aprotinin occurs by a caspase-dependent mechanism through the mitochondrial pathway.

Fig. 3. Aprotinin Inhibits Apoptosis via Mitochondrial Pathway and 250KIU/ml is an Optimal Dose for Protection.

Fig. 3

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(a,b) Caspase 3/7 and 9 activities after 24hrs SD in primary cortical neuron culture. (c-e) Images after 24hrs SD in a neuron culture without Aprotinin. (f) Percentage of caspase3+ cells at 24hrs after SD with Aprotinin at the concentration of 0, 100, 250, 500, and 1000KIU/ml. *p < 0.05, **p < 0.01, ***p < 0.001 vs. Aprotinin 0KIU/ml by ANOVA with Bonferroni comparisons. Data are shown as mean ± SEM (n = 6-8).

Finally we assessed the anti-apoptotic effect of aprotinin from 100KIU/ml up to 1000KIU/ml, in order to identify the optimal dose for adjunctive neuroprotection. Using immuhistochemistry, 100 and 250KIU/ml aprotinin dose-dependently decreased caspase3+ cell percentages after 24hrs SD (Fig. 3c-f). Aprotinin at a concentration of 500 and 1000KIU/ml also reduced the percentage compared with No aprotinin; however, these doses did not provide significant reduction as compared with 250KIU/ml (Fig. 3c). This suggests that approximately 200-250KIU/ml is an optimal dose of aprotinin for adjunctive protection during cardiac surgery.

Discussion

This study is the first to describe neuroprotective potentials of three anti-fibrinolytic agents utilized during cardiac surgery against glutamatergic excitotoxicity in mixed cortical cell culture models. We then demonstrated the anti-apoptotic effects of aprotinin using various analyses in primary neuronal cell culture system. We finally identified the optimal dose for neuroprotection in the clinical application during cardiac surgery.

Recently the role of anti-fibrinolytic agents on postoperative neurological events has come into increasing focus. High-dose TXA has been identified as an independent predictor of early seizures after CPB in clinical studies.17, 18 A retrospective data analysis has also demonstated that even moderate TXA doses are associated with a doubled rate of convulsive seizures and in-hospital mortality.19 Consistent with these findings, preclinical study by Lecker et al identified that the reduction in function of glycine receptors caused by TXA leads to disinhibition and proconvulsive effects.20 Since their results and those of others21 determined that TXA is a competitive antagonist of GABAA receptors, it is suggested that TXA inhibition of GABAA receptors increases network excitability.20 EACA is also a competitive antagonist of glycine receptors, but not aprotinin.20 In the present study, we investigated the neuroprotective potential of antifibrinolytic agents against glutamatergic excitotoxicity that is a major mechanism of ischemic neuronal cell death. We used an in vitro culture model, but did not study electrophysiological activity. However, we recently developed an ex vivo rodent brain slice model replicating specific brain conditions of deep hypothermic circulatory arrest including ischemia-reperfusion/reoxygenation under hypothermia.22 Receptor expression and function on neurons and glial cells are dynamically changing during development;23 however, there is little information regarding the effects of pharmacological reagents including anti-fibrinolytics during cardiac surgery on electrophysiological activity in the developing brain. Rodent brain slice models have been used widely for electrophysiological studies and for the development of pharmacologic therapy. Thus the model will be useful for the study of optimal anti-fibrinolytic agents in pediatric cardiac surgery. We believe our future studies using the in vitro neuronal cell culture model, ex vivo brain slice model, and in vivo large animal model will provide novel insights regarding cellular and molecular mechanisms underlying neurological deficits after cardiac surgery, and allow for the design of targeted therapies and conditions which will minimize the risk of neurodevelopmental deficits in CHD patients.

In 2007, the US Food and Drug Administration suggested to the Bayer Corporation that it should suspend marketing of aprotinin following a retrospective propensity match review by Mangano et al and preliminary analysis of a prospective trial in Canada (BART trial).24, 25 The Bayer Corporation responded by voluntarily removing all remaining stocks of aprotinin from the US market in 2008.25 However, there has been accumulating evidence of the deleterious effects of alternative hemostatic agents such as Factor VII and lysine analogues as well as the disadvantages of massive transfusion, suggesting that the protective effects of aprotinin outweighed the risks involved.26-28 Based on subsequent analysis of the BART trial, Health Canada concluded in 2011 that the trial could not be reliably used to assess the benefit–risk balance of aprotinin as a result of study weaknesses.29 The European Medicines Agency also released a statement in 2012, indicating that the benefits of aprotinin outweigh its risks in patients and recommended to the EU that the suspension of the licence for aprotinin in this context be lifted.29

In addition to functioning as an important haemostatic agent, it has been suggested that aprotinin inhibits nitric oxide synthase activity30 and the proinflammatory activation of endothelial cells.31 Using both a piglet model involving cardiopulmonary bypass as well as a neuronal cell culture model, we have obtained abundant evidence for the neuroprotective effects of aprotinin.5, 6, 32-34 Together with our present result, it can be inferred that aprotinin is the preferred anti-fibrinolytic agent during cardiac surgery, in particular for patients at high risk of procedure related brain injury such as those undergoing circulatory arrest. It is likely that aprotinin will be reintroduced into the US through a distributor other than the Bayer Corporation. This will allow clinical trials to explore the neuroprotective effects of aprotinin as studied in this report.

Conclusions

Aprotinin at 200-250 KIU/ml exerts maximal protective effects on primary cortical neurons cultured in vitro. This effect might result from caspase-dependent and/or independent pathways. In conjunction with our previous studies, aprotinin is a multipotent serine protease inhibitor which can protect the brain during cardiac surgery.

Acknowledgments

We thank Dr David Zurakowski for assistance with statistical analysis. We are thankful to Dr Olivier Nicole for assistance in the preparation of the cell culture model.

Funded by NIH R01HL060922 (R.A.J) and R01HL104173 (R.A.J)

Footnotes

Disclosure: Authors have nothing to disclose with regard to commercial support.

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