Identification of Fibrinogen as a Key Anti-apoptotic Factor in Human Fresh Frozen Plasma for Protecting Endothelial Cells in vitro

Abstract

Resuscitation with human fresh frozen plasma (FFP) in hemorrhagic shock (HS) patients is associated with improved clinical outcomes. Our group has demonstrated that the beneficial effect of FFP is due to its blockade on endothelial hyperpermeability, thereby improving vascular barrier function. The current study aimed to investigate HS-induced endothelial cell apoptosis, a potential major contributor to the endothelial hyperpermeability, and to determine the effect and the key components/factors of FFP on protecting endothelial cells from apoptosis. We first measured and demonstrated an increase in apoptotic endothelial microparticles (CD146⁺AnnexinV⁺) in patients in shock compared to normal subjects, indicating the induction of endothelial cell activation and apoptosis in shock patients. We then transfused HS rats with FFP and showed that FFP blocked HS-induced endothelial cell apoptosis in gut tissue. To identify the anti-apoptotic factors in FFP, we utilized high performance liquid chromatography, fractionated FFP, and screened the fractions in vitro for the anti-apoptotic effects. We selected the most effective fractions, performed mass spectrometry, and identified fibrinogen as a potent anti-apoptotic factor. Taken together, our findings suggest that HS-induced endothelial apoptosis may constitute a major mechanism underlying the vascular hyperpermeability. Furthermore, the identified anti-apoptotic factor fibrinogen may contribute to the beneficial effects of FFP resuscitation, and therefore, may have therapeutic potential for HS.

INTRODUCTION

Trauma is the leading cause of death in the United States and worldwide (1, 2). Up to 30% of trauma deaths are due to uncontrolled blood loss or hemorrhagic shock (HS) (3, 4). One of the primary clinical manifestations of HS is the disruption of the vascular barrier, which leads to microvascular hyperpermeability in vital organs (5). Activation of endothelial apoptotic signaling may contribute to microvascular hyperpermeability (5–9).

Our group and others applied damage control resuscitation as rapid hemorrhage control through early administration of blood products in a balanced ratio (1:1:1 for units of plasma to platelets to red blood cells) and found that the balanced ratio transfusion reduced mortality (10–13). Our group has further demonstrated in pre-clinical studies that administration of fresh frozen plasma (FFP) protected hemorrhage-induced damage of vital organs by reducing vascular permeability in animals and blocked hypoxia-induced endothelial hyperpermeability in vitro (14–18). Moreover, we showed that human plasma protects against hypoxia and serum starvation-induced endothelial apoptosis in vitro (19). However, it is largely unknown which components/factors are responsible for the anti-apoptotic effect in FFP. FFP consists of over 3600 proteins and is the most complex human-derived proteome (20–23). Identification of the effective components in FFP responsible for the anti-apoptotic activity may lead to novel therapeutic development for resuscitation in HS patients.

In this study, we demonstrated increased apoptotic endothelial microparticles in plasma of patients with severe injuries and in shock compared with healthy controls. We also showed in HS rat model that FFP resuscitation inhibited HS-induced endothelial cell apoptosis. We further verified the anti-apoptotic effect of FFP on endothelial cells in vitro. Moreover, we took a proteomic approach and explored the effective anti-apoptotic components/factors in FFP, identified and validated fibrinogen as a potent anti-apoptotic factor.

MATERIALS AND METHODS

Reagents

Human FFPs were obtained from the Gulf Coast Regional Blood Center (Houston, TX, USA). FFPs pooled from three donors were used for the animal experiments and most of the in vitro studies. The FFP used for fractionation was from a single donor. The native form of human fibrinogen was obtained from Novus Biologicals (Centennial, CO, USA). Recombinant human proteins of ApoE3 and CD14 were obtained from R&D Systems (Minneapolis, MN, USA).

Human subject study

The human subject study was approved by The University of Texas Health Science Center at Houston (UTHSC-H) Committee for the Protection of Human Subjects through waiver of informed consent. As described previously (18), we measured plasma adiponectin levels from severely injured patients in hemorrhagic shock and demonstrated significantly lower adiponectin levels at the time of admission to the emergency department (ED) compared with healthy donors, and a slight increase of adiponectin levels upon admission to the intensive care unit (ICU), but not significant compared to admission to the ED. The same set of severely injured patients in hemorrhagic shock was used for the study, with a systolic blood pressure <90 mmHg and/or a base deficit of >5 meq/mL, and received at least one unit of blood after ED arrival but before admission to ICU. Patients’ demographics, laboratory values, and outcomes were obtained from patient records. Blood samples were obtained upon admission to the ED and subsequently upon arrival to the ICU at Memorial Hermann Hospital in Houston, Texas, a Level I trauma center. Blood samples were also obtained from healthy subjects following consent. After collection, blood samples were centrifuged, and plasma aliquots were prepared and stored at −80°C for subsequent analysis

Endothelial microparticle measurement from the human plasma

Endothelial microparticles (EcMPs) were measured by multicolor flow cytometry as previously described (24, 25). Briefly, the human plasma samples were stained with an endothelial cell marker CD146-PE and an apoptotic cell marker AnnexinV-FITC. The double positive CD146⁺AnnexinV⁺ EcMPs were calculated and expressed as apoptotic EcMPs.

Rat HS model and resuscitation

All animal procedures were approved by the Animal Welfare Committee at the UTHSC-H. All animal experiments were performed according to the guidelines of the Animal Welfare Act and the Guide for Care and Use of Laboratory Animals from the NIH. A Sprague-Dawley rat HS model was used as previously described (14). Briefly, the rats were anesthetized, intubated, and ventilated. Tygon catheters were placed into the abdominal aorta through the femoral artery to record arterial blood pressure and heart rate, as well as into the femoral vein for resuscitation. HS was induced by withdrawing blood from the femoral vein until mean arterial blood pressure (MAP) stabilized at 25 mmHg. One hour after HS, animals were randomly assigned to HS alone without resuscitation or HS+FFP resuscitation. FFP was infused after HS to a volume equal to the blood lost (range of 2 mL/100 g body weight) (14). Sham group received the same procedure except blood withdrawal or resuscitation. Three hours after FFP resuscitation, the rats were euthanized and the gut was harvested for further analysis.

Evaluation of gut injury and the vascular cell apoptosis from the HS rats

Gut injury was assessed on H&E stained paraffin sections using the Chiu score for histopathological evaluation (26). Briefly, the gut injury was scored by a reviewer blinded to the treatment using a semi-quantitative grading system based on the mucosal and submucosal damage (0 represents normal, 1 to 5 represents injury from mild to severe).

Detection of vascular apoptosis in the gut tissue was performed on the paraffin sections using VasoTACS in situ apoptosis detection kit (Trevigen, Inc., Gaithersburg, MD) according to the manufacturer’s instruction.

Endothelial cell culture and treatment

Human pulmonary microvascular endothelial cells (HPMECs) were purchased from PromoCell (Heidelberg, Germany) and cultured in endothelial cell growth medium (Promocell) as recommended by the manufacturer. The cells were used within passage 10 for all experiments.

The cells were seeded in 96-well plates (2 × 10⁴/well) overnight, and then serum starved (0.1% FBS) for 4 hours. The cells were pretreated with FFP, FFP fractions, or identified factors for 30 min, followed by staurosporine (STS, 1 μM) treatment for 3 hours for apoptosis analysis.

Apoptosis analysis

At the end of the treatment, the cells were lysed for Caspase-3/7 activity assay using a luminescent Caspase-Glo 3/7 Assay kit (Promega, Madison, WI) according to the manufacturer’s instructions. Apoptosis was also measured by quantification of DNA fragmentation using a cell death detection ELISA assay (Roche Molecular Biochemicals, Indianapolis, IN) according to the manufacturer’s instruction and as previously described (27–29).

FFP fractionation by high performance liquid chromatography

Citrated FFP was fractionated through high performance liquid chromatography (HPLC) by size exclusion, using a sepharose CL-4B column at a controlled flow rate of 1.2 ml/min. A total of 107 fractions were obtained. We adopted a two-step screening process to identify active molecule(s) because the large numbers of fractions generated using HPLC would be very difficult to test individually in a consistent manner. For the first step, we grouped 54 odd number fractions into 6 large pools, each containing 9 odd numbered fractions. The remaining aliquots of the 54 odd number fractions and the 53 even number fractions were kept individually and stored at −80°C until experiments. Testing pooled fractions in the first screen step allowed us to narrow down which pooled fraction contained the anti-apoptotic activity. Once the endothelial protective activity was detected in a specific pool, odd numbered fractions in that pool were examined individually and verified in the corresponding even numbered fractions to identify the active factor(s).

Mass spectrometry analysis

Fractions 92, 94, and 106 were selected for mass spectrometry (MS) for identifying the active factor(s). They were subjected to acetone precipitation at −20°C overnight. After centrifugation (12,000 g × 5 min), the pellets were reduced with 30 μl of 6 M urea, 20 mM DTT in 150 mM Tris HCl, pH 8.0, at 37°C for 40 min, then alkylated with 40 mM iodacetamide in the dark for 30 min. The reaction mixture was diluted 10-fold using 50 mM Tris-HCl pH 8.0, followed by overnight digestion at 37°C with trypsin (1:20 of enzyme trypsin vs protein substrate). Digestions were terminated by adding an equal volume of 2% formic acid, and then desalted using Waters Oasis HLB 1 ml reverse phase cartridge according to the vendor’s procedure. Eluates were dried via vacuum centrifugation.

An aliquot of the tryptic digest (in 2 % acetonitrile/0.1% formic acid in water) was analyzed by LC/MS/MS on an Orbitrap Fusion™ Tribrid™ mass spectrometer (Thermo Scientific™) interfaced with a Dionex UltiMate 3000 Binary RSLCnano System. Peptides were separated onto an Acclaim™ PepMap™ C₁₈ column (75μm ID × 15 cm, 2 μm) at a flow rate of 300 nl/min. Gradient conditions were: 3%−22% B for 120 min; 22%−35% B for 10min; 35%−90% B for 10 min; 90% B held for 10 min (solvent A, 0.1 % formic acid in water; solvent B, 0.1% formic acid in acetonitrile). The peptides were analyzed using a data-dependent acquisition method. Orbitrap Fusion was operated with measurement of FTMS1 at resolution 120,000 FWHM, scan range 350–1500 m/z, AGC target 2E5, and maximum injection time of 50 ms. During a maximum 3 second cycle time, the ITMS2 spectra were collected at rapid scan rate mode, with CID NCE 35, 1.6 m/z isolation window, AGC target 1E4, maximum injection time of 35 ms, and dynamic exclusion was employed for 60 seconds.

The raw data files were processed using Thermo Scientific™ Proteome Discoverer™ software version 1.4, spectra were searched against the Uniprot-Homo sapiens database using the Mascot search engine. Search results were trimmed to a 1% false discovery rate (FDR) using Percolator. For the trypsin, up to two missed cleavages were allowed. MS tolerance was set to 10 ppm; MS/MS tolerance 0.6 Da. Carbamidomethylation on cysteine residues was used as fixed modification; oxidation of methionine, as well as phosphorylation of serine, threonine and tyrosine, were set as variable modifications.

Statistical analysis

Data are expressed as mean ± SEM. Statistical significance between multiple groups was determined by one-way ANOVA followed by Holm-Sidak test using SigmaPlot 11.0 (Systat Software, Chicago, Illinois). p values less than 0.05 are considered significant.

RESULTS

HS induces EC apoptosis in patients

HS-induced endothelial cell apoptosis studies have mainly been performed in animal HS models (9, 30, 31), largely due to the unavailability of human tissue samples from HS patients. Microparticles (MPs) are small cellular membrane fragments shed from activated or apoptotic blood cells and endothelial cells (32). Apoptotic endothelial MPs (EcMPs) can be readily detected from human plasma using an endothelial marker, CD146, and an apoptosis marker, AnnexinV, as we previously reported (24). In this study, to evaluate endothelial apoptosis in HS patients, we performed flow cytometry to identify apoptotic EcMPs stained by both CD146 and AnnexinV. We studied twenty patients with severe injuries in shock (median injury severity score 34 (29, 36.5), base deficit 7 (7, 10)) that survived to the ICU and had a 30% mortality rate. Compared with healthy controls (48±5, n=12), patients in shock had higher numbers of apoptotic EcMPs (double positive for CD146⁺AnnexinV⁺) at the time of admission to ED (425±115, n=20, p<0.05) and persisted into the ICU (421±106). These data suggest that HS induces endothelial apoptosis in human patients who are severely injured. Although all patients received FFP, FFP resuscitation may protect endothelial cells from further injury, but may not eliminate already formed EcMPs.

FFP protects against HS-induced gut injury and vascular cell apoptosis in a rat model

We used a rat HS model as previously described (14) to evaluate the injury of the gut, one of the sensitive organs to HS damage, using a semi-quantitative Chiu scoring system (26). As shown in Fig. 1A and 1B, an increase in the Chiu scores was observed in HS group compared with sham controls (3.00±0.57 vs 0.13±0.13, p<0.05), and Chiu scores were reduced in HS+FFP group compared with HS group (1.00±0.11 vs 3.00±0/57, p<0.05). To investigate whether HS induces endothelial cell apoptosis, we stained the gut tissue sections with VasoTACS in situ apoptosis detection kit. As demonstrated in Fig. 1C and 1D, we observed an increased number of stained apoptotic cells in HS group compared with sham group (527±26 vs 56±19, p<0.05), and the number of apoptotic cells was reduced in HS+FFP group compared with HS group (89±12 vs 527±26, p<0.05). Taken together, our results indicate that HS induces gut endothelial apoptosis in rats, and FFP protects against HS-induced gut endothelial apoptosis.

Fig. 1. FFP protects against HS-induced gut injury and vascular cell apoptosis in a rat model.

Rat gut tissue samples were collected from the indicated groups: sham, HS, HS+FFP. A.Representative images of H&E staining and B. Quantification of gut injury by Chiu scores. C.Representative images of VasoTACS in situ staining of apoptotic endothelial cells (as pointed by arrows) and D.Quantification of apoptotic endothelial cells. n=4–7 rats/group. Data are expressed as mean ± SEM. * p<0.05 compared with Sham. # p<0.05 compared with HS alone.

Identification of the effective anti-apoptotic fractions from FFP

We have previously shown that human plasma protects against hypoxia and serum starvation induced endothelial apoptosis in vitro (19). In the present study, we used a potent apoptosis inducer, staurosporine (STS) (33, 34), and screened multiple FFP pools and fractions for the anti-apoptotic function. We first confirmed the apoptosis inducing effect of STS and the anti-apoptotic effect of FFP by measuring Caspase 3/7 activity and DNA fragmentation. As demonstrated in Fig. 2A, STS induced a >2.5-fold increase of Caspase 3/7 activity compared with vehicle control (p<0.05). FFP pretreatment at 10% in culture medium abolished STS-induced Caspase 3/7 activity (p<0.05). A similar pattern was observed in the DNA fragmentation measurement (Fig. 2B). These results are consistent with our previous study using hypoxia and serum starvation induced apoptosis (19). Since the Caspase3/7 assay is designed for use with multiwell-plate formats, ideal for high-throughput screening, validated by using several apoptosis-inducing agents including staurosporine, resulting in a rapid and sensitive Caspase-3/7 activity assay, we used the Caspase-3/7 activity assay in the following experiments to efficiently screen multiple FFP fractions and factors.

Fig. 2. FFP blocks STS-induced apoptosis in HPMECs in vitro.

HPMECs were seeded and cultured in the growth medium overnight. The cells were starved with 0.1% FBS for 4 hours, pretreated for 30min with human fresh frozen plasma (FFP, 10%), followed by staurosporine (STS) treatment (1 μM) for 3 hours. A. Caspase 3/7 activity and B. DNA fragmentation were measured. n=4 wells/group. Data are expressed as mean ± SEM. *p<0.05 compared with vehicle control group. ^# p<0.05 compared with STS group.

We pooled FFP fractions as illustrated in Table 1, and screened their effects on STS-induced apoptosis in HPMECs. The pools 3, 4, 5, and 6 revealed anti-apoptotic effects, and significant differences were observed across pools 3 to 6 (Fig. 3A). We went on to further screen individual fractions from the two most effective pools 5 and 6, and identified several fractions with anti-apoptotic effects (Fig. 3B and 3C).

Table 1.

FFP fractions and the pools.

Pools	Fractions
P1	1	3	5	7	9	11	13	15	17
P2	19	21	23	25	27	29	31	33	35
P3	37	39	41	43	45	47	49	51	53
P4	55	57	59	61	63	65	67	69	71
P5	73	75	77	79	81	83	85	87	89
P6	91	93	95	97	99	101	103	105	107

Fig. 3. Identification of the effective FFP pools and fractions for protecting HPMECs from apoptosis induction.

HPMECs were cultured as described in Fig. 2. A. Cells were pretreated for 30min with the indicated pools (10%), followed by STS (1 μM) treatment for 3 hours. FFP (10%) was used as control. B and C. Cells were pretreated with the indicated individual fractions (10%) that were included in pool 5 or pool 6, respectively. Caspase 3/7 activity was measured. n=4 wells/group. Data are expressed as mean ± SEM. *p<0.05 compared with STS group. #p<0.05 across pools 3 to 6.

Identification of the anti-apoptotic factors from the FFP fractions

To identify the anti-apoptotic factors in these fractions, we chose the even fractions 92 and 94, which were close to the effective odd number fractions 91 and 93 and had the least freezing and thawing cycles, and performed mass spectrometry (MS). The even fraction 106 was used as a negative control (Fig. 3C).

MS analysis identified a total of 140, 136, and 116 proteins in fractions 92, 94, and 106, respectively, with some overlap among the fractions (Fig. 4). We focused on the 51 proteins overlapping only between fractions 92 and 94, and performed UniProt database (http://www.uniprot.org) searching for their anti-apoptotic functions. We identified 3 candidate proteins: fibrinogen, apolipoprotein (Apo) E3, and monocyte differentiation antigen CD14.

Fig. 4. Venn diagram showing the mass spectrometry (MS) outcome and screening of the anti-apoptotic factors.

Total proteins identified by MS from each of the fractions, with the overlapping proteins among the fractions, are presented in the Venn diagram. The 51 proteins that overlap between fractions 92 and 94 were searched for anti-apoptotic function against the database. The indicated three proteins were identified.

We took the same approach as described in Fig. 3 and validated the effect of fibrinogen, ApoE3, and CD14 on STS-induced apoptosis. As demonstrated in Fig. 5, fibrinogen pretreatment inhibited STS-induced Caspase 3/7 activity at concentrations of 625 and 1250 μg/ml (p<0.05) in HPMECs. ApoE3 and CD14 pretreatment did not inhibit STS-induced Caspase 3/7 activity in HPMECs. Thus, we have identified and validated fibrinogen as a potent anti-apoptotic factor in FFP.

Fig. 5. Validation of the identified proteins from MS for the anti-apoptotic function in vitro.

HPMECs were cultured as described in Fig. 2. Cells were pretreated for 30min with the indicated proteins at different concentrations, followed by STS (1 μM) treatment for 3 hours. FFP (10%) was used as control. Concentrations of the proteins used for pretreatment: Fibrinogen at 125, 625, and 1250 μg/ml, ApoE3 at 5, 50, and 200 μg/ml, and CD14 at 0.5, 2.5, and 10 μg/ml. Caspase 3/7 activity was measured. n=4 wells/group. Data are expressed as mean ± SEM. *p<0.05 compared with STS group.

DISCUSSION

The results of this study demonstrate that HS induces endothelial apoptosis in human patients and animal models, which may contribute to the vascular hyperpemeability in HS and can be mitigated by FFP resuscitation. Via a proteomic approach, we have identified several candidate proteins from FFP and validated fibrinogen as a potent anti-apoptotic factor.

One of the primary clinical manifestations of HS is the disruption of the vascular barrier, which leads to microvascular hyperpermeability in vital organs and contributes to the morbidity and mortality of shock (5, 35). In addition to the vascular barrier regulation, vascular cell apoptosis may compose a major pathological mechanism underlying the microvascular hyperpermeability. However, most studies in HS-induced endothelial cell apoptosis were performed in animal HS models (9, 30, 31). To establish clinical relevance of the studies on endothelial apoptosis in HS patients, we measured the number of endothelial microparticles that bear an apoptotic marker (CD146⁺AnnexinV⁺), a surrogate of endothelial apoptosis, in HS patient plasma. We found a significant increase of apoptotic EcMPs in HS patients compared with normal human subjects, suggesting that HS induces endothelial apoptosis. The results of HS-induced endothelial apoptosis in the rat model support the findings from the human study.

FFP resuscitation after HS is clinically beneficial to HS patients (10–13), and reduces vascular hyperpermeability in HS animal models and in cultured endothelial cells in vitro (14–18). In this study, we demonstrated that FFP can also inhibit HS-induced endothelial apoptosis in rat gut tissue (Fig. 1) and STS-induced endothelial apoptosis in cultured endothelial cells in vitro (Fig. 2).

Plasma is a complex biologic material that contains thousands of proteins covering a myriad of physiological and pathological functions (20–23, 36). To identify which components/factors in FFP have the anti-apoptotic function, we first applied HPLC, collected 107 fractions, and screened the fractions for their anti-apoptotic function in vitro. We then chose the most effective fractions and performed mass spectrometry analysis. Through database searching, several candidate proteins were identified including fibrinogen, ApoE3, and CD14.

Fibrinogen is coagulation factor I, abundant in normal plasma with concentration of 2.0–4.5 mg/ml (37). Fibrinogen is a homodimer with a molecular size of 340 kDa, consisting of two subunits, an alpha and a beta chain. Fibrinogen is secreted primarily by hepatocytes and plays a central role in clot formation. By interacting with intercellular adhesion molecule 1, fibrinogen promotes endothelial cell survival, thus has an anti-apoptotic effect (38). Clinical transfusion of fibrinogen to HS patients is being evaluated for its potential effect on survival after injury (39–42). In this study, we have validated the anti-apoptotic function of fibrinogen in HPMECs in vitro, and demonstrated that the anti-apoptotic effect of fibrinogen is significant at 0.625 and 1.25 mg/ml. The effect of 1.25 mg/ml is potent, although below its biological concentration (2.5–4 mg/ml), and comparable to the effect of 10% FFP (Fig. 5). Thus, our findings further support the clinical use of fibrinogen and FFP transfusion in HS patients for the benefit of providing coagulation factors, as well as diminishing endothelial injury, thus improving vascular function.

ApoE is a 34 kDa protein with four major isoforms, ApoE1, ApoE2, ApoE3, and ApoE4. ApoE3 is the most common form. The concentration of ApoE in normal plasma is 30–70 μg/ml (43). ApoE is important in lipid metabolism. It can also stimulate neurite outgrowth and antagonize ApoE4-induced neuronal cell apoptosis (44). CD14 is a 55 kDa cell surface glycoprotein that is preferentially expressed on monocytes/macrophages. CD14 exists in two forms, a membrane bound and a soluble form. The membrane bound CD14 is important for the recognition and clearance of apoptotic cells (45). It also acts as a coreceptor for Toll-like receptors that bind bacterial lipopolysaccharide, triggering inflammatory responses (46). The soluble CD14 either appears after shedding off the membrane or is directly secreted by the liver and monocytes. The soluble CD14 is present in normal plasma with concentration of 2–6 μg/ml (47). It protects chronic lymphocytic leukemia cells from apoptosis (48). However, in our study, pretreatment with ApoE3 or CD14 did not suppress STS-induced Caspase 3/7 activity in the human pulmonary microvascular endothelial cells. Thus, the contrasting results from our studies using an endothelial model, and from the literature using neurites and lymphocytes, may be due to the use of different cell models.

Several limitations are considered in this study. First, a single donor was used for plasma fractionation and the following mass spectrometry. However, the anti-apoptotic effect of the identified fractions from the single donor plasma are comparable with 10% FFP that consists of 3 donors. In addition, a mass spectrometry profile derived from the plasma of a single healthy donor was able to represent healthy control in comparison with that in trauma patients (21). Second, fraction 106 may not be an ideal negative control since it is next to fraction 105 that still demonstrated a noticeably, although weaker, anti-apoptotic effect. We have only focused on the 51 proteins that overlap between fractions 92 and 94 in the current study. Thus, the selection of the pools and fractions for apoptosis screening and MS analysis, and the selected screening of the proteins following MS analysis could not cover all potential effective fractions/factors. Third, pretreatment with fibrinogen and other proteins for the in vitro studies may not mimic the exact in vivo resuscitation regimen. Therefore, the more relevant in vivo studies are crucial and planned as future studies for validating the findings from the in vitro studies presented currently. Furthermore, the protective effects of FFP resuscitation are multifactorial. Several factors that can reduce HS-induced endothelial hyperpermeability have been reported by our group (16, 18), including a recent in vitro study regarding the effect of fibrinogen on protecting endothelial against barrier dysfunction (49). Thus, our current study has revealed another aspect of fibrinogen, its anti-apoptotic property.

Overall, our findings provide further evidence to support HS-induced endothelial apoptosis as a pathophysiological mechanism underlying microvascular hyperpermeability in HS, and support the anti-apoptotic role of FFP resuscitation in protecting endothelial cells in HS. The identified anti-apoptotic fractions derived from FFP and the anti-apoptotic factor fibrinogen provide the opportunity for the development of mechanism-based therapeutic interventions as surrogate to FFP for treatment of trauma/HS patients.