Platelets are vital for the regulation of hemostasis. Platelets produced in the bone marrow circulate, and are destroyed in the spleen after ∼7 days in circulation. Normal platelet counts in circulation average around 150,000 to 400,000/μL. Platelets are transfused to prevent or treat hemorrhage in hypoproliferative thrombocytopenia, and in a variety of settings such as cardiopulmonary bypass-related platelet consumption and dysfunction, drug-induced platelet inhibition, and severe trauma. Prophylactic transfusion is generally initiated when the platelet count drops below 10,000/μL, though certain clinical scenarios can prompt prophylaxis at higher counts, such as 50,000/uL for surgery or 100,000/uL for neuraxial surgery (1). Current US blood-banking practice dictates that platelets be stored at 22°C (RT) in an incubator with gentle agitation for not more than 5 days, as it has been established that platelets stored under these conditions survive for up to 7 days after transfusion in vivo(2). The increase in platelet in-vivo survival comes at the cost of viability and functionality. As the hemostatic and metabolic functions of RT-stored platelets decline significantly there is also an increased risk of bacterial contamination (3). A viable alternative is the storage of platelets at 4°C (4C) or cold storage. Though cold storage reduces the circulation time by half, it also reduces risk of bacterial growth, preserves in-vitro measures of hemostatic function, effectively reverses bleeding time prolongation due to aspirin, and reduces bleeding in thrombocytopenic patients (4–6).
We recently showed that apheresis platelets (AP) stored in cold, compared with room temperature, have more viable metabolic characteristics, perform better in functional tests, form stronger clots, and release fewer inflammatory mediators (7). In our study, we also found that heightened functional responses of refrigerated platelets may be attributed to the increased activation status of these platelets, as measured by CD62P expression and phosphatidylserine (PS) exposure. Consistent with what others have shown, our study indicated that though refrigerated platelets are viable and functional, they are in an activated or “primed” state. Thus, the transfusion of an activated platelet product may carry the risk of inappropriate aggregation and thrombotic complications.
Under normal conditions, healthy endothelium secretes nitric oxide (NO) and prostacyclin (PGI2) into the bloodstream which keep platelets circulating in a quiescent state (8). NO release from endothelial cells via endothelial nitric oxide synthase is regulated by physiological levels of shear stresses associated with blood flow. NO passively diffuses through the plasma membrane to bind intracellular soluble guanylyl cyclase (sGC) (9). This binding causes upregulation of cyclic guanosine monophosphate (cGMP) and activation of protein kinase G (PKG). PKG activation leads to phosphorylation of inositol-1, 4, 5-triphosphate, blocks agonist-induced rises in intracellular calcium (10), and subsequently decreases platelet aggregation (11, 12). Prostacyclin, in contrast, acts primarily on platelet surface prostacyclin (IP) receptors. Binding of prostacyclin to an IP receptor leads to activation of adenylyl cyclase, formation of cyclic adenosine monophosphate (cAMP), and phosphorylation of protein kinase A (PKA) (8). PKA phosphorylation, like PKG phosphorylation, leads to reduced intracellular calcium levels and inhibits platelet functions such as activation and aggregation.
Platelet storage studies have focused largely on the pre-activation mechanisms that lead to attenuated response to agonists, which are collectively known as platelet storage lesions (3, 13). However, very little is known about the effect of storage on the response of platelets to physiological inhibitors of activation, and the associated signaling pathways. Kobsar et al. (14) showed that RT-stored apheresis platelet concentrates exhibit increased cGMP accumulation after 6 days of storage, but no studies to date have been conducted using 4C-stored platelets. In this study, we treated apheresis platelets stored at both RT and 4C with the NO donor, s-nitrosoglutathione (GSNO), and PGI2 before testing platelet-specific function. Both GSNO and PGI2 are well characterized inhibitors of platelet aggregation and adhesion in in vitro settings (15, 16). We hypothesized that the physiologic inhibitors NO and PGI2 would inhibit 4C-stored platelet adhesion and aggregation response, demonstrating that refrigerated platelets respond to physiologic controls.
MATERIALS AND METHODS
GSNO, bovine serum albumin (BSA), and phosphate buffered saline (PBS) were purchased from Sigma Aldrich (St. Louis, Mo). The platelet agonists, adenosine diphosphate (ADP), collagen, and thrombin receptor activating peptide-6 (TRAP) were obtained from Diapharma, West Chester, OH, and prepared in PBS. PGI2, cGMP EIA kits, and cAMP EIA kits were purchased from Cayman Chemicals, Ann Arbor, MI. FITC-conjugated bovine lactadherin and the anti-human sheep polyclonal antibody, FITC-conjugated anti-human Factor V (PAHFV-S), were obtained from Haematologic Technologies, Inc, Essex Junction, VT. The anti-human monoclonal mouse antibody, APC-conjugated anti-CD62P (P-selectin, clone AK4), was purchased from BD Biosciences, San Jose, CA. Calcein-AM, Fluo-4 AM, and DiOC6 were all obtained from Invitrogen Life Technologies, Carslbad, CA.
Platelet storage and handling
Single AP units were collected from healthy donors (n = 4) in acid-citrate-dextrose (ACD)-plasma and 10-mL aliquots were sterilely transferred into 15 mL BCSI mini-bags (Seattle, WA) for storage as previously described (7). Mini-bags were stored for 5 days either at RT (RT-PLT) in a Food and Drug Administration (FDA)-approved platelet incubator with agitation or in a walk-in refrigerator (4C-PLT) without agitation. Platelets were assayed on the day of collection (Day 1, FR-PLT) and Day 5. 4°C-stored samples were allowed to come to RT for 30 min prior to testing. Where applicable, platelet poor plasma (PPP) was obtained by centrifugation of AP for 10 min at 3,000 × g. In some studies, whole blood was collected in ACD vacutainer tubes and spun at 3,500 × g for 10 min to obtain RBCs. Washed platelets were obtained by centrifugation of AP at 900 × g for 10 min to obtain a pellet, which was resuspended in Ca+2/Mg+2-free Tyrode's Buffer (NaCl 137 mM, KCl 2.7 mM, MgCl 2 mM, NaH2PO4 0.42 mM, Glucose 5 mM, and HEPES 10 mM). Prior to testing, some samples were treated to with ABT-737 (30 μM) for 2 h at 37°C, A23187 (10 μM) for 10 min at 22°C, or TRAP (20 μM) for 10 min at 22°C to serve as positive controls for activation.
Calibrated automated thrombogram
Thrombin generation in platelet samples diluted in autologous plasma (150,000/μL) was measured using the calibrated automated thrombogram (CAT) method (Thrombinoscope BV, Maastricht, The Netherlands) as previously described (17). Inhibited samples were treated with 100 nM PGI2 for 2 min before testing commenced. Thrombin generation was detected using a Fluoroskan Ascent fluorometer (Thermo Fisher Scientific, Waltham, MA) for 50 min. Thrombin generation curves, lag times, endogenous thrombin potentials (ETP), and peak thrombin concentrations were analyzed using Thrombinoscope software version 220.127.116.112. Lag time corresponded to the clotting time, peak height was the maximum thrombin concentration achieved, and ETP was the area under the curve or total amount of thrombin generated.
Surface-expressed factor V (FV)
Activated platelet surface expression of FV was assessed using a polyclonal antibody against human FV followed by flow cytometry. Prior to immunostaining, platelets were incubated with Human TruStain FcX (BioLegend, San Diego, CA) for 5 min at RT, followed by PE-conjugated FV antibody at RT for 20 min in the dark. The samples were diluted to 1 mL with PBS and analyzed by flow cytometry (FACS Canto).
Intracellular free calcium
Platelet intracellular free calcium was determined as previously described (18). Briefly, 50 uL platelet samples (untreated, ABT-737, and A23187; 300,000/μL) were incubated with 10 uL Fluo-4 AM (2 μM final concentration) for 20 min at RT in the dark. Upon incubation, samples were then diluted to 500 uL with Tyrode's Buffer and analyzed with flow cytometry as previously described. Mean channel fluorescence (MCF) was divided by mean forward scatter (FSC) to generate intracellular free Ca2+ as previously described (18).
Dense granule release of ATP
Platelet dense granule secretion was assessed by the measurement of ATP release as previously described (19) using a luciferin-luciferase reagent or a Chrono-Lume reagent (DuPont, Wilmington, DE). Secretion was measured using a lumi-aggregometer (Chrono-log Corp, Havertown, PA).
Platelet activation markers
Platelets treated with inhibitors (10 nM PGI2 for 2 min or 500 μM GSNO for 5 min) or Tyrode's buffer (untreated) were activated with TRAP (20 μM) and ADP (50 μM) and assessed for the activation markers CD62P (P-Selectin) and phosphatidylserine (PS) surface expression (lactadherin binding) using flow cytometry (7).
96-well plate aggregation
Platelets were assessed for aggregation response using the 96-well plate method as previously described (20). 90 μL of AP diluted with PPP to a concentration of 300,000/μL was added to a 96-well plate containing 10 μL of the platelet agonists ADP (1, 10, and 20 μM final concentrations), collagen (2, 5, and 10 μg/mL final concentrations), and TRAP (25, 50, and 100 μM final concentrations). In some experiments, PRP was treated with 10 nM PGI2 for 2 min or 50 μM GSNO for 5 min prior to addition of PRP to the 96-well plate. The plate was then immediately placed in a microplate reader and absorbance was determined at 650 nm every 10 s for 10 min between vigorous shaking at 37°C.
Multiplate impedance aggregation
Impedance aggregometry was performed as previously described (7). 300 μL of AP diluted with PPP to a concentration of 300,000/μL was briefly incubated with 300 μL of 3 mM NaCl2/CaCl2 solution (Verum Diagnostica, Munich, Bavaria). Upon a brief incubation period, 20 μL of agonist (32 μM TRAP, 3.2 μg/mL Collagen, and 6.4 μM ADP final concentrations) was added. All tests were carried out for 12 min, and area under the curve (AUC) was reported. PGI2 and GSNO were added to some samples prior to assay as described above.
Flow-based adhesion studies
Bioflux plates (Fluxion Biosciences, San Francisco, CA) were coated with 100 μg/mL type I collagen from equine tendon (Helena Laboratories, Beaumont, TX) for 1 h at 22°C. Channels were then rinsed with PBS, and blocked with 0.5% BSA in PBS for 15 min. Platelets were stained with calcein-AM (10 μM final concentration) for 30 min at 22°C before resuspending them in plasma at a concentration of 300,000/μL and 40% hematocrit with RBCs, and perfused through collagen-coated wells at a wall shear rate of 720/s. Some platelets were treated with PGI2 and GSNO prior to perfusion, as described above. Fluorescence microscopy was used to visualize thrombus formation and pictures were acquired every 30 s for 6 min. Fluorescence intensity units (FIU) and Surface Coverage were estimated using Bioflux Montage MetaMorph software (Fluxion Biosciences).
cGMP and cAMP accumulation were determined in washed platelets according to previously established methods (21). Washed platelets (300,000/μL) were incubated with 500 μM GSNO for cGMP measurements or with 10 nM PGI2 for cAMP measurements or in Tyrode's Buffer without the inhibitor, for 5 min. To stop the reaction, the platelets were immediately placed in liquid nitrogen. Platelet pellets were then hydrolyzed with 15 μL of 3.3 M HCl, the lysates were then centrifuged for 10 min at 600×g, and the supernatants were assayed for cGMP and cAMP, respectively, using EIA kits according to the manufacturer protocols.
Inhibition data were analyzed by two-way ANOVA with a post-hoc Tukey's test for multiple comparisons. All other data were analyzed by one-way ANOVA and Student t test. Significance was set at P < 0.05. Data in graphs are mean ± SEM. Data management was achieved using Microsoft Excel (Microsoft Corp., Redmond, WA) and statistical analyses were performed using GraphPad Prism 6 (GraphPad Software Inc, San Diego, CA).
Accelerated thrombin generation in 4C-stored platelets is due to increased FV
To evaluate the thrombin generation capability of stored platelets, we chose to utilize the CAT assay (Fig. 1). Testing thrombin generation has been widely considered a standard function test of the coagulation system with physiological relevance. In particular, the calibrated automated thrombogram assay is unique in that it has the ability to generate almost 100 thrombin generation curves at once and is more sensitive than other coagulation tests in terms of distinguishing hypo- and hyper-coagulability (22). A typical thrombogram produced from the CAT assay resembles a bell-shaped curve and contains several distinguishing features including a lagtime in which the concentration of thrombin increases from 0 nM to 5 nM, and a peak representing the maximum thrombin concentration generated (23). In this study, we chose to report the lagtime, peak height, and endogenous thrombin potential, a representation of the conceivable amount of enzymatic work from thrombin over the course of its lifetime (24).
A significant decrease in lag time was seen in cold-stored platelets compared with fresh platelets (Fig. 1B), while a significant increase in peak height was observed in cold-stored platelets compared with both fresh and RT-stored platelets (FR-PLT vs. 4C-PLT, P < 0.005; RT-PLT vs. 4C-PLT, P < 0.05, Fig. 1C). However, no significant differences were seen between any groups in terms of endogenous thrombin potential.
To determine the cause of the rapid generation of thrombin observed in 4C-PLT, we assessed the presence of coagulation factor expression, focusing our attention on factor V (FV). Platelet α-granules contain FV that can be released upon platelet activation (25). This platelet-released FV has been shown to play a large role in the promotion and maintenance of hemostasis at sites of injury (25). We observed 3-fold higher levels of FV expression (% total and GMFI) in cold-stored platelets compared with both fresh (%: P < 0.001; GMFI: P < 0.01) and 1.5-fold higher than RT platelets (%: P = 0.03; GMFI: P = 0.01) after 5 days of storage (Fig. 1, E and F).
Intracellular free calcium and dense granule release are increased in 4C platelets
Upon activation, calcium is released from primarily endoplasmic reticulum stores and the dense tubular system into the cytosol. Calcium plays a large role in regulating platelet activation, aggregation, and adhesion (26). We assessed intracellular free calcium in fresh and stored platelets, and discovered that cold-stored platelets showed significantly higher levels than both fresh and RT (Fig. 2A).
Platelet dense granules contain adenosine triphosphate (ATP) that is released upon platelet activation. We observed a 2-fold decrease in ATP release in both RT (P = 0.021) and 4C-stored platelets (P
= 0.016) compared with fresh (Fig. 2B). However, no difference was observed between RT and 4C.
Intracellular cGMP/cAMP levels are maintained during storage
Having established that stored platelets despite are activated, we tested the response of stored platelets to physiologic inhibitors, PGI2 and GSNO. The binding of PGI2 to platelet IP receptors activates adenylate cyclase resulting in an increase in the intracellular cAMP levels (9). GSNO is a NO donor with low cytotoxicity and potent anti-platelet effects (27). The NO released by GSNO passes through the platelet membrane, activates sGC, and leads to an increase in the levels of intracellular cGMP (10). We used ELISA to measure the concentration of cAMP and cGMP in fresh platelets and platelets stored at RT or 4C (Fig. 3). When the platelets were not treated with the inhibitors, we did not observe any significant differences in cAMP or cGMP concentration between fresh and stored samples. When platelets were treated with PGI2, cAMP levels rose by more than 2-fold from basal levels in fresh platelets (P < 0.05) and cold-stored platelets (P < 0.05), but not in RT-stored platelets. GSNO treatment caused a significant 4-fold increase of cGMP in all storage conditions (FR-PLT P < 0.01; RT-PLT P < 0.001; 4C-PLT P < 0.0001). These data suggest that platelet inhibitory machinery is preserved during storage at both RT and 4C.
Cold-stored platelet activation, but not thrombin generation, is inhibited by prostacyclin and nitric oxide
We examined whether thrombin generation can be attenuated with physiologic inhibitor, PGI2. As expected, prostacyclin treatment significantly reduced peak height (P < 0.01) and endogenous thrombin potential in fresh platelets, but had no effect on lagtime (Table 1). Contrastingly, treatment with PGI2 had no effect on platelets stored at either RT or 4C.
Next, we evaluated the effect of the inhibitors PGI2 and GSNO on platelet activation in response to TRAP by measuring the surface expression of two established markers of activation: exposure of phosphatidylserine (PS) and upregulation of P-selectin released by the α-granules. The concentrations of GSNO and PGI2 used throughout this study were determined from dose response of fresh platelets (Figure, Supplemental Digital Content 1, at http://links.lww.com/SHK/A334 [Supplemental Figure 1: Inhibitor log dose response-curves. Normalized data for (A) PGI2 and (B) GSNO. Data are expressed as mean ± SEM for n = 2 donors.]). We observed that, consistent with our previous work (7), storage significantly increases both CD62P expression levels and PS exposure on platelets stored at both RT and 4C (Fig. 4). Interestingly, TRAP treatment activates stored platelets even further, reaching levels comparable to TRAP-activated fresh platelets (Fig. 4, A–C). TRAP-induced platelet activation in both fresh and stored platelets is abrogated upon treatment with PGI2 and GSNO. Our data suggest that stored platelets can still be activated by an agonist and the activation can be inhibited by physiologic inhibitors.
Cold-stored platelet aggregation is inhibited by prostacyclin and nitric oxide
We evaluated the response of inhibitor-treated platelets to low, normal, and supra-maximal concentrations of the agonists ADP, collagen, and TRAP using 96-well plate aggregometry and impedance aggregometry, as described in the Methods section (Fig. 5). As expected, the aggregation response of untreated, fresh, and stored platelets increased with the increase in agonist concentration (Fig. 5, A1–C3). Both PGI2 and GSNO treatment attenuated the aggregation response of fresh and stored platelets to all concentrations of ADP and collagen tested in this study (Fig. 5, A1–B3). In contrast, while PGI2 attenuated platelet aggregation at all concentrations of TRAP in both fresh and stored samples in the 96-well assay, GSNO was effective in blocking aggregation at high concentrations of TRAP only in fresh platelets but not in RT or 4C platelets (Fig. 5, C1–C3); however, the degree of GSNO inhibition in stored samples was comparable to that seen in fresh. The inability of GSNO to block aggregation of stored platelets at supra-maximal concentrations of TRAP can likely be attributed to a combination of the high agonist potency, the amount of platelets that are pre-activated due to storage, and the low inhibitor potency of GSNO (28). While GSNO managed to significantly inhibit Multiplate impedance aggregation of 4C platelets (Fig. 5, F) to TRAP, the level of inhibition was significantly different from the levels of inhibition observed in both fresh and RT-treated platelets. This difference may be due to the physical means of measurement between optical and impedance aggregometry—optical aggregometry detects changes in light transmission resulting from platelet shape change and aggregation, whereas Multiplate aggregometry records changes in resistance from platelet adherence to the electrodes (29).
Cold-stored platelets are more adhesive than RT-stored platelets
We evaluated the effect of inhibitors on platelet adhesion to injured subendothelium. To simulate blood flow over injured endothelium, platelets in reconstituted blood were perfused at an arterial shear rate through channels coated with collagen containing trace amounts of vWF. The channels were coated with BSA to prevent non-specific adhesion of platelets (Fig. 6). We observed that the adhesion and aggregation of untreated fresh and cold-stored platelets were comparable as estimated from surface coverage and total fluorescence, respectively (Fig. 6). Platelet adhesion and aggregation in both fresh and cold-stored platelets were abrogated upon treatment with PGI2 and GSNO. RT-stored platelets did not adhere to the collagen surface at all, suggesting platelet dysfunction and possibly degradation of plasma and coagulation factors RT storage (30).
We have previously shown that refrigerated platelets though primed, are responsive to agonist stimulation (7). In this study, we show that refrigerated platelets generate thrombin more rapidly, presumably due to increased PS exposure and FV on their surface. We also show that 4C platelets, although primed, maintain their responsiveness to physiologic inhibitors, nitric oxide, and prostacyclin, as these inhibitors are effective in curtailing the activation, adhesion and aggregation responses to agonists. Together, our data suggest that refrigerated platelets are a hemostatically better product with intact intracellular regulation.
Activated platelets express negatively charged phosphatidylserine on their outer surface, wherein tenase and prothrombinase complexes assemble to generate thrombin. It has been reported previously that platelet activation during storage generates microparticles or extracellular vesicles, which are potently procoagulant due to high levels PS exposed on their surface, and these play a very important role in hemostasis (31, 32). Thus, the shortened lag times, i.e., faster thrombin generation, observed in both RT and 4C-stored platelets, compared with fresh platelets, is likely attributed to increased surface PS expression and increased microparticles. However, the differences in lag time and peak thrombin observed between RT and 4C samples may be due to the significantly higher FV expression in 4C-stored platelets than in RT-stored platelets since FV, and its activated form (FVa), is crucial for the formation of the prothrombinase complex and is responsible for raising factor Xa's activity, and hence thrombin generation, by more than 5-fold (26). Our data also suggest that increased platelet activation during both cold and RT storage is due to an increase in intracellular calcium concentration, which may arise either from the release of intracellular calcium stores or from the entry of extracellular calcium through the plasma membrane (33).
Endothelial cells lining the vasculature release NO and PGI2, which are potent inhibitors of platelet activation. A pivotal step in the inhibition of platelets by PGI2 and NO is the increase in intracellular cAMP and cGMP levels, respectively. The basal levels of cAMP and cGMP in fresh and stored platelets measured in this work were comparable to published values (34). Although a rise in cGMP, but not in cAMP, levels was reported in apheresis platelets stored at RT for 6 days (14), we did not observe any substantial differences in basal cGMP or cAMP after 5 days of storage. PGI2-treatment caused significant increases in cAMP for fresh and cold-stored platelets but only modestly in RT-stored platelets, possibly as a result of defective intracellular signaling induced by the platelet storage lesion (3, 14). However, it has been shown that even slight rises in cAMP are enough to inhibit platelet aggregation to arachidonic acid (35). As expected, GSNO treatment led to significant increases in cGMP accumulation in all storage groups.
We analyzed the effect of increases in cAMP and cGMP levels on the activation and aggregation of platelets in response to agonists. We chose the platelet agonists ADP, collagen, and TRAP as each of these agonists activates platelets through distinct receptors, namely, P2Y1, P2Y12, and GPVI, respectively. Fresh, RT-, and cold-stored platelets are activated by agonists, and both PGI2 and GSNO significantly inhibited platelet activation, though not to the degree of baseline levels. This result suggests that the activation during storage is, at least to some extent, reversible and does not represent a terminal activation state.
Increased activation may lead to thrombosis when transfusing platelets in vivo. As a way to assess the safety of fresh and stored platelets we utilized the physiologic inhibitors, PGI2 and GSNO, in several functional assays. As expected (10, 15, 16), PGI2 and GSNO were effective at inhibiting fresh platelet aggregation. Remarkably, the addition of PGI2 and GSNO suppressed the agonist-induced aggregation of cold-stored platelets to levels comparable to that of fresh platelets for all agonists in both light transmission and impedance aggregometry. Consistent with our previous data (7), RT-stored platelets failed to respond to ADP and collagen and only responded to TRAP, a strong agonist. It is relevant to note that aggregometry is widely regarded as the clinical gold standard for diagnosing platelet hemostatic dysfunction in vivo. The data presented here further underscore the hemostatic similarity between cold-stored and fresh platelets.
In a trauma scenario, a platelet product must be able to quickly respond and repair the damaged endothelium. This complex biophysical process involves initial adhesion of platelets to subendothelial matrix proteins including collagen and VWF, aggregation, and subsequent recruitment of platelets from flowing blood to the growing thrombus by dense granule release or production of soluble agonists. Our data showed that cold-stored platelets adhere and form thrombi similar to that of fresh platelets, and these responses can be attenuated by the use of endothelial inhibitors. Of note, adhesion of RT-stored platelets was very poor despite the addition of fresh RBCs used in the assay suggesting that during trauma, hemostatically weak RT-stored platelets may provide, at best, only minimal benefit. In every functional test performed in this study, and in our previous work, RT-stored platelets showed a significant decline in platelet function compared with both fresh and 4C-stored platelets. While RT-stored platelets express activation markers and generate copious thrombin, they are not truly “functional.” Cold storage of platelets may result in a better hemostatic product compared with RT and better in vivo and in vitro function may in turn decrease demands on a limited inventory by reducing transfusion volumes (4–7, 30).
In summary, we showed that cold-stored platelets, despite showing higher activation levels than fresh platelets, still maintain intact intracellular control circuitry. The activation, adhesion and aggregation responses of cold-stored platelets may be switched on and off by physiologic concentrations of agonists and inhibitors, respectively. Further, these responses are comparable to fresh platelets, and are more sensitive and robust than RT-stored platelets. Thus, this study on in vitro safety of cold-stored platelets, together with our previous work on in vitro efficacy, further strengthens the case to consider clinical study of cold-stored platelets for therapeutic transfusion.
The authors would like to acknowledge the US Army Institute of Surgical Research blood bank for collection of apheresis units.
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