Pathogen‐reduced plasma, cryoprecipitate reduced for therapeutic plasma exchange

Abstract

Background

Therapeutic plasma exchange (TPE) for thrombotic thrombocytopenic purpura (TTP) and auto‐immune disorders involves repeated patient exposure to allogenic plasma with the risk of transfusion‐transmitted infection (TTI). Amotosalen‐UVA Pathogen Reduction technology is FDA approved to manufacture pathogen‐reduced plasma, cryoprecipitate reduced (PRPCR), a form of cryoprecipitate poor plasma (CPP) with potentially improved TPE outcomes and reduced TTI risk.

Methods

PRPCR was manufactured from pathogen‐reduced (PR) plasma. Thrombin generation, fibrinogen, Factors II, V, VII, VIII, IX, X, XI, XIII, VWF, ADAMTS13, Protein C, Protein S, α‐2 plasmin inhibitor (α‐2 PI), IgG, IgM, and IgA were measured. Microfluidic chamber assays at variable shear rates characterized PRPCR‐mediated platelet adhesion and aggregation.

Results

Compared to PR plasma, fibrinogen, Factor VIII, and VWF levels were depleted in PRPCR. Factors II, V, VII, IX, X, XI, XIII, thrombin generation, Protein C, Protein S, α‐2 PI, ADAMTS13, and immunoglobulins were conserved. At low wall shear rates (300 s⁻¹) PRPCR supported platelet adhesion. Perfusion of plasma‐free blood containing PRPCR flowed over immobilized VWF binding peptide (100 μg/mL) and showed absence of platelet adhesion. Perfusion of plasma‐free blood containing PRPCR flowed over immobilized collagen (200 μg/mL) at high wall shear rate (1500 s⁻¹) and demonstrated no platelet thrombus formation.

Conclusions

PRPCR retained hemostatic capacity, anti‐thrombotic proteins, and ADAMTS13, but collagen induced platelet aggregation was negligible at high shear due to depletion of functional high molecular weight VWF. PRPCR is a CPP option for TPE with reduced platelet‐mediated thrombotic risk and TTI risk, but with retention of plasma hemostatic capacity and immunoglobulins.

Abbreviations

CPP

cryoprecipitate poor plasma

ETP

endogenous thrombin potential

FFP

fresh frozen plasma

HSA

human serum albumin

PR

pathogen‐reduced

PRPCR

pathogen‐reduced plasma, cryoprecipitate reduced

RIPA

ristocetin‐induced platelet aggregation

TPE

therapeutic plasma exchange

TTI

transfusion‐transmitted infection

TTP

thrombotic thrombocytopenic purpura

VWF

von Willebrand factor

α‐2 PI

α‐2 plasmin inhibitor

1. INTRODUCTION

Therapeutic plasma exchange (TPE) is an effective treatment for patients with thrombotic thrombocytopenic purpura (TTP) and other autoimmune disorders. ¹ TPE has reduced the mortality of TTP from >80% to <10%. ² In TTP, the primary objective of TPE is to correct the deficiency of the von Willebrand factor (VWF)‐cleaving protease ADAMTS13 and to remove anti‐ADAMTS13 antibodies. ³ , ⁴ However, effective TPE often requires episodic repeat treatments, resulting in repeated exposure to large volumes of allogeneic donor plasma and increased risk of transfusion‐transmitted infection (TTI). ⁵ , ⁶ , ⁷ Bacterial contamination of plasma is generally considered low risk due to frozen storage; however, bacteria can contaminate donor plasma and remain viable in frozen blood products. ⁸ , ⁹ In vulnerable populations such as patients with autoimmune disorders, even low bacterial loads may pose an infectious risk. To mitigate TTI risk, including unrecognized or emerging pathogens for which specific donor testing may not be available, pathogen inactivation technologies using photochemical or solvent/detergent methods have been implemented. ¹⁰ , ¹¹ , ¹² , ¹³

Pathogen‐reduced (PR) platelets have been widely adopted in the United States and Europe to reduce infectious risks from bacteria and emerging pathogens, and analogous PR approaches have been applied to products used for TPE. Replacement fluid options for TPE include 5% albumin, fresh frozen plasma (FFP), conventional cryoprecipitate‐poor plasma (CPP), solvent/detergent‐treated plasma or PR plasma. ¹⁴ , ¹⁵ In TTP, guidelines recommend replacement with a product containing ADAMTS13. ¹ Accordingly, multiple pathogen‐inactivated plasma derivatives have been used for TPE. ¹⁶ , ¹⁷ , ¹⁸ , ¹⁹ CPP is an alternative to plasma because ADAMTS13 is conserved ¹⁶ , ²⁰ , ²¹ , ²² , ²³ ; with the potential advantage of reduced procoagulant activity and decreased platelet activation due to depletion of FVIII, fibrinogen, and VWF while retaining immunoglobulins, and sufficient levels of other coagulation factors to support hemostasis during TPE. ²³ , ²⁴ By potentially limiting platelet activation and thrombotic risk during recovery, particularly when platelet counts rise rapidly, CPP could reduce the need for adjunctive antiplatelet or anticoagulant therapy recommended by some clinicians. ²⁵ Because the pathophysiology of TTP involves ultra‐large VWF multimers driving platelet‐rich microthrombi formation in the setting of acquired ADAMTS13 deficiency, CPP depleted of ultra‐large VWF multimers has been investigated as a potentially superior replacement fluid. ²⁶ , ²⁷ Recently, Mafra et al. reported a meta‐analysis showing CPP significantly reduced mortality in comparison to fresh frozen plasma for treatment of TTP. ²⁸ While both plasma and CPP are indicated for TPE of TTP, conventional plasma and PR plasma have been preferred modalities for TPE, and PR CPP was previously not available. ¹⁷ , ²⁹

In 2020, FDA approved pathogen‐reduced plasma, cryoprecipitate reduced (PRPCR), a form of PR CPP, with a primary indication for TPE in TTP and a secondary indication for coagulation factor replacement excluding fibrinogen, FVIII, FXIII, and VWF. ³⁰ As PRPCR is a plasma derivative, it is important to ensure that it contains the elements required for effective TPE, without inducing excessive platelet adhesion, activation and aggregation, which could result in thrombotic adverse events. The current study characterizes the constituents and function of PRPCR with respect to platelet adhesion and aggregation under variable shear flow conditions. A dynamic study of platelet functionality using microfluidic assays simulates the physiology of rheologic events in vessels to evaluate the potential of critical proteins to support or facilitate platelet adhesion, activation and aggregation under physiologically relevant shear conditions.

2. METHODS

2.1. Manufacture of pathogen‐reduced plasma, cryoprecipitate reduced (PRPCR) and pathogen reduced cryoprecipitate fibrinogen complex (PRCFC)

PR plasma, PRPCR and PRCFC were prepared by licensed blood establishments produced as commercial components per manufacturer's instructions (Central California Blood Center, Fresno, CA), and were shipped frozen on dry ice by World Courier with temperature monitoring. Blood collection from volunteer donors (group A or O) was in accordance with FDA regulations and the AABB Circular of Information. PRPCR (Cerus Corporation, Concord, CA) and Pathogen‐Reduced Cryoprecipitated Fibrinogen Complex (PRCFC, INTERCEPT Fibrinogen Complex‐IFC, Cerus Corporation, Concord, CA) were prepared from plasma (pooled from ABO‐identical donors) separated from four whole blood plasma units anticoagulated with acid citrate dextrose. Plasma (~650 mL) was prepared using amotosalen (150 μM) and long wavelength ultraviolet light (UVA, 320–400 nm) in a 3 J/cm² treatment. ³⁰ , ³¹ , ³² Amotosalen intercalates into helical regions of DNA and RNA in dark equilibrium and illumination with UVA light (320–400 nm) results in the formation of covalent irreversible amotosalen nucleic acid adducts inhibiting pathogen replication and infectious potential. ³¹ Reduction of residual amotosalen was achieved by filtration through an integral compound absorption device (Figure S1). Following preparation of PR plasma, the treated plasma was transferred into a single container and frozen (−18°C to −25°C). Within 30 days of freezing, the frozen PR plasma was thawed at 4°C–6°C to produce a PR cryoprecipitated fibrinogen complex (PRCFC) enriched with fibrinogen, Factor VIII, VWF, Factor XIII, and Fibronectin. The cryoprecipitate container was centrifuged to separate the PRCFC from the PRPCR supernatant and both were stored frozen at −18°C to −25°C with 1 year expiration. ³⁰ For the microfluidic experiments, fibrinogen concentrations and not VWF concentrations were adjusted as PRPCR contains very low levels of VWF. The fibrinogen concentration of PRCFC and PRPCR was adjusted to 0.5 mg/mL in PBS when coating chambers or in Tyrode's albumin buffer when used to prepare plasma‐free blood. The fibrinogen concentration of 0.5 mg/mL preserved platelet aggregation in response to ADP indicative of sufficient fibrinogen to support platelet aggregation. ³³ After dilution based on a fibrinogen concentration of 0.5 mg/mL, the final VWF activity level in PRCFC was 0.28 IU/mL. The diluted PRCFC served as a positive control for assessment of fibrinogen and VWF function in the microfluidic assays. Undiluted PRPCR contained negligible VWF activity (Table 1) that was not changed significantly by dilution based on fibrinogen content.

TABLE 1.

Comparative coagulation factor activities in PRPCR and PR plasma.

Coagulation factor activity	PRPCR	PR plasma
Thrombin generation‐ETP (nM min)	1156 ± 208	1581 ± 154
Fibrinogen (mg/mL)	1.47 ± 0.15	2.28 ± 0.49
Factor II (IU/mL)	0.77 ± 0.07	0.89 ± 0.12
Factor V (IU/mL)	0.66 ± 0.12	0.86 ± 0.17
Factor VII (IU/mL)	0.83 ± 0.21	0.77 ± 0.23
Factor VIII (IU/mL)	0.15 ± 0.05	0.92 ± 0.35
Factor IX (IU/mL)	1.00 ± 0.19	1.00 ± 0.25
Factor X (IU/mL)	0.86 ± 0.13	0.94 ± 0.19
Factor XI (IU/mL)	1.02 ± 0.29	0.92 ± 0.21
Factor XIII antigen (IU/mL)	1.05 ± 0.20	1.32 ± 0.15
Von Willebrand factor RIPA (IU/mL)	0.10 ± 0.00	0.95 ± 0.38

Note: The values for coagulation factor activity in PR plasma (n = 64) were determined in FDA registration studies and published in the product package insert for FDA registration. ³⁷ The values for coagulation factor content in PRPCR (n = 60) were determined in FDA registration studies and published in the product package insert. ³⁰ Mean values (SD) for thrombin generation ETP and coagulation factor activities in PRPCR and PR Plasma are summarized from the published product package inserts derived from separately conducted studies and thus, no statistical comparisons are made. Generally established reference range values for pathogen reduced (PR) donor plasma coagulation factor activities are 0.5–1.5 IU/mL and for fibrinogen 200–400 mg/dL. Reference range data for conventional plasma are published in the AABB Circular of Information for the Use of Human Blood and Blood Components. ⁴⁹ There are no reported reference ranges for coagulation factor activity in conventional cryoprecipitate reduced plasma. Reported reference ranges for thrombin generation expressed as endogenous thrombin potential (ETP) are 1424–3024 nM min. ³²

2.2. Assay of functional coagulation factors and immunoglobulin

PRPCR and PR plasma were characterized by Cerus Corporation using validated assays. Fibrinogen, Factor II, Factor V, Factor VII, Factor VIII:C, Factor IX, Factor X, and Factor XI were assayed using one stage clotting assays with serial dilutions and reference standards (Diagnostica Stago, Inc., Parsippany, NJ). Thrombin generation, expressed as endogenous thrombin potential (ETP), was assayed by calibrated automated thrombogram with 5 picomolar tissue factor (Fluroskan Ascant Fluorometer, Thermo Fisher Scientific, Corston, Bath, UK). Factor XIII antigen was measured by K‐Assay (Stago Compact Analyzer, Diagnostica Stago, Inc., Parsippany, NJ). VWF activity was measured by ristocetin‐induced platelet aggregation (RIPA) cofactor assay with serial dilutions (Chrono‐Log Aggregometer, Havertown, PA). ADAMTS13 was measured by immunoassay (Quantikine Human ADAMTS13, R&D Systems, Minneapolis, MN). Protein C and Protein S were measured by functional clotting assays (Diagnostica Stago, Inc., Parsippany, NJ). Αlpha‐2 plasmin inhibitor was measured by immunoassay (Euroimmun, Lubeck, Germany). IgG, IgM, and IgA were measured by Becton Dickinson Cytometric Bead Array Human Immunoglobulin Flex Set System (Becton Dickinson Biosciences, Milpitas, CA) and SDS PAGE assay (Prothya Biosolutions, Brussels, Belgium).

2.3. Preparation of plasma free washed platelets and red blood cells

Blood samples for use in microfluidic assays were collected from healthy volunteer blood donors, who gave written, informed consent according to institutional guidelines (Établissement Français du Sang [EFS], Strasbourg, France). Washed human platelets were prepared as described previously. ³⁴ Platelets were isolated from human whole blood anticoagulated with ACD by differential centrifugation and washed twice at 37°C in Tyrode's buffer (137 mM NaCl, 2 mM KCl, 12 mM NaHCO₃, 0.3 mM NaH₂PO₄, 1 mM MgCl₂, 2 mM CaCl₂, 5.5 mM glucose, 5 mM HEPES, pH 7.3, 295 mOsm L⁻¹) containing 0.35% purified human serum albumin (HSA) and 0.5 μM prostaglandin I2 (PGI₂). Finally, platelets were suspended in Tyrode's buffer containing 0.35% HSA and 0.02 U mL⁻¹ of the adenine nucleotide scavenger apyrase and were incubated at 37°C throughout all experiments.

To prepare a suspension of washed red blood cells (RBCs), fresh hirudin‐anticoagulated (100 U/mL) blood obtained from healthy donors was centrifuged at 250 × g for 16 min. The platelet‐rich plasma and leukocyte layers were removed, and the RBCs were washed twice in plain Tyrode's buffer (137 mmol/L NaCl, 2 mmol/L KCl, 12 mmol/L NaHCO₃, 0.3 mmol/L NaH₂PO₄, pH 7.3) and finally suspended in Tyrode's buffer (137 mmol/L NaCl, 2 mmol/L KCl, 12 mmol/L NaHCO₃, 0.3 mmol/L NaH₂PO₄, 1 mmol/L MgCl₂, 2 mmol/L CaCl₂, 5.5 mmol/L glucose, 5 mmol/LHEPES, pH 7.3, 295 mOsmol), containing 0.35% human serum albumin (HSA), 0.02 U/mL of the adenine nucleotides scavenger apyrase and 100 U/mL hirudin. Finally, RBCs were pelleted by a 3000 g centrifugation for 4 min and used for microfluidic assays during the following 90 min.

2.4. In vitro microfluidic assays with variable shear forces

Microfluidic flow chambers were prepared according to the method previously described. ³⁵ The channels were coated with PRCFC or PRPCR (0.5 mg/mL of fibrinogen, diluted in PBS) for 2 h at room temperature, or with a VWF‐binding peptide (100 μg/mL, provided by Professor Richard Farndale) ³⁶ overnight at 4°C, or with type I fibrillary collagen (200 μg/mL, Takeda, France) for 1 h at room temperature; and blocked with PBS containing 1% fatty acid‐free human serum albumin (HSA) for 30 min at room temperature. A programmable syringe pump (PHD 2000, Harvard Apparatus) was used to perfuse blood through the coated channels. Hirudin‐anticoagulated (525 ATU/mL) whole blood obtained from healthy donors was perfused at 37°C in microfluidic chambers containing immobilized PRCFC or PRPCR at 300 s⁻¹ for 5 min. For inhibition studies, the blood was incubated with abciximab (40 μg/mL), an αIIbβ3 antagonist, for 10 min at 37°C before perfusion, or with caplacizumab (10 μg/mL), a VWF antagonist added to the blood just prior perfusion. Reconstituted plasma‐free blood composed of washed platelets (1 × 10⁵/μL), washed RBCs (40% hematocrit), PRCFC, PRPCR (0.5 mg/mL of fibrinogen diluted in Tyrode's buffer) or Tyrode's buffer and hirudin (100 U/mL) was perfused at 1500 s⁻¹ over VWF‐binding peptide for 3 min or at 3000 s⁻¹ over collagen for 5 min. PRCFC was used as a positive control in microfluidic assays; and HSA or Tyrode's buffer was used as a negative control in microfluidic assays. The average fibrinogen concentration of PRCFC for these experiments was 8.21 ± 0.84 mg/mL and the average VWF activity of PRCFC was 4.55 ± 0.43 IU/mL. To increase the sensitivity to detect changes of platelet interactions with fibrinogen and VWF, the fibrinogen concentration of PRCFC and PRPCR was adjusted to 0.5 mg/mL resulting in an average VWF activity concentration of 0.28 IU/mL for PRCFC. These concentrations are below the reference values of whole blood from healthy donors, in which fibrinogen concentration is 2 to 4 mg/mL and VWF concentration is 0.5–2.0 IU/mL.

Platelet adhesion on immobilized PRCFC, PRPCR and VWF‐binding peptide was monitored in real time with an inverted Leica DMI8 microscope (Leica Microsystems) using a 40×, 0.6 numerical aperture oil objective, a Leica K5 sCMOS 4.2 MP camera and a differential interference contrast (DIC) technique. For thrombus formation on collagen‐coated microfluidic chamber, platelets were labeled with DIOC₆ (1 μmol/L, Molecular Probes, UK) for 10 min at 37°C before perfusion. After excitation with a 488 nm argon‐ion laser, fluorescence emission was measured in real time using a confocal Leica SP8 inverted microscope with a resonant scanner and a 40x oil objective. Series of optical sections in xyz from the base to the peak of the thrombi were recorded over time. The Images were stacked and analyzed with ImageJ software (National Institutes of Health, Bethesda, MD, USA) to determine the number of adherent platelets and the thrombus volume. The total analyzed surface area was >50,000 μm².

2.5. Statistical analysis

Data for coagulation factor activities, inhibitor proteins, and immunoglobulins were characterized by mean and standard deviations (SD) with reference to normal range values. The characteristics of PRPCR and PR Plasma are provided to characterize these components, and they were not analyzed for statistical differences because these products were prepared from different plasma collections over time and mean values represent the content of these products. Statistical analysis of microfluidic assays was performed using GraphPad software (Prism 9.2.0). Data were reported as mean ± standard error of the mean (SEM). After establishment of the parametric distribution of the data using the Shapiro–Wilk test, data were analyzed using one‐way ANOVA followed by Bonferroni post hoc test. A p‐value of <0.05 was considered statistically significant, and other p‐values denoted by * as follows: ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

3. RESULTS

3.1. Content of plasma proteins in PRPCR and pathogen‐reduced plasma

Plasma protein activities in PR Plasma (n = 64) and PRPCR (n = 60) were extracted from previously published FDA registration studies and product package inserts, ³⁰ , ³⁷ and are used here for comparative purposes. PRPCR, a PR CPP, contains decreased levels of fibrinogen, Factor VIII, and von Willebrand factor compared to PR plasma, which is commonly used for TPE (Table 1). ¹⁷ Factor XIII antigen was lower in PRPCR than in PR plasma, but it was retained within the normal reference range. Coagulation factors II, V, VII, IX, X, and XI were retained at therapeutic levels in PRPCR (Table 1). Thrombin generation, an integrated parameter of hemostatic capacity ³⁸ expressed as endogenous thrombin potential (ETP), was reduced in PRPCR compared to PR plasma, but with residual thrombin generation activity (Figures S2 and S3, Table 1). ADAMTS 13 and anti‐thrombotic protein activities were retained in PRPCR at similar levels to PR plasma (Table 2). In addition, PRPCR (n = 12) retained immunoglobulin concentrations similar to those of PR plasma (n = 6) without qualitative changes (Table 2, Figure 1).

TABLE 2.

Activity of ADAMTS13, anti‐thrombotic proteins, and immunoglobulin content in PRPCR and PR plasma.

Factor activity	PRPCR	PR plasma
ADAMTS 13 functional (IU/mL)	1.32 ± 1.8	0.90 ± 1.6
Protein C (IU/mL)	0.99 ± 0.14	0.79 ± 0.18
Protein S (IU/mL)	0.83 ± 0.10	0.97 ± 0.23
α‐2 Plasmin inhibitor (IU/mL)	0.86 ± 0.13	0.76 ± 0.07
IgG (g/L)	9.6 ± 4.5	7.22 ± 0.92
IgA (g/L)	2.30 ± 1.20	1.49 ± 0.65
IgM (g/L)	0.70 ± 0.50	0.64 ± 0.21

Note: Mean values (SD) for ADAMTS 13, Protein C, Protein S, and α‐2 plasmin inhibitor in PRPCR (n = 60) and PR Plasma (n = 64) are summarized from product package inserts derived from separate studies for PRPCR and PR Plasma referenced in Table 1, and no statistical comparisons are made. For ADAMTS 13, Protein C, Protein S, and α‐2 plasmin inhibitor, reference range values for conventional plasma are 0.5–1.5 IU/mL. Reference range values for Protein C and Protein S are described in the AABB Circular of Information for the Use of Human Blood and Blood Components. Reference range data for ADAMTS13 and α‐2 plasmin inhibitor are not reported in the AABB Circular of Information for the Use of Human Blood and Blood Components. Values for IgG, IgA, and IgM in PRPCR (n = 12) were measured after 30 days of frozen storage (−18°C to −25°C) prior to analysis. Values for IgG, IgA, and IgM in PR plasma (n = 6) were measured in components after 3 months of frozen storage (−18°C to −25°C). PRPCR and PR plasma were evaluated by SDS‐PAGE analysis for qualitative changes performed as described in Figure 1. These studies demonstrated no qualitative changes in IgG.

FIGURE 1.

Reducing PAGE Analysis of Reference Plasma and PRPCR. Reducing SDS‐PAGE analysis was used to detect qualitative changes in immunoglobulins. Sample 141122 is reference plasma; samples 080323 and 140323 are PRPCR. The two large bands are reduced heavy and light IgG chains with molecular weights of 50 and 25 kDa, respectively. The other bands are due to partially reduced IgG. The absence of a smear at the bottom of the gel indicates that no IgG polymers were formed. Amotosalen treatment did not affect IgG based on molecular weight analyses. The samples shown are representative of six PR plasma and 12 PRPCR components.

3.2. Functional properties of PRPCR to support platelet adhesion under shear flow

We used microfluidic assays to characterize the functional properties of PRPCR in mediating platelet adhesion and aggregation under variable wall shear flow. Flow chambers were coated with PRPCR or PRCFC, using equalized fibrinogen levels (0.50 mg/mL). PRCFC was used as a positive control rather than plasma because it is the cryoprecipitate generated during PRPCR manufacturing from PR plasma and is therefore enriched in essential proteins for platelet adhesion, including fibrinogen and VWF. ³³ The average VWF activity in the diluted PRPCR was <0.10 IU/mL and 0.28 IU/mL in the diluted PRCFC positive control. At large artery wall shear rates (300 s⁻¹), perfusion with whole blood for 5 min resulted in efficient platelet adhesion to surfaces coated with PRCFC, PRPCR, but not to surfaces coated with HSA (Figure 2A). At similar fibrinogen concentrations (0.5 mg/mL), PRPCR demonstrated a slightly reduced level of platelet adhesion (PRCFC: 32718 ± 4928 plts/mm² vs. PRPCR: 23182 ± 5841 plts/mm²; n = 7 blood donors), but adhesion was not statistically different from PRCFC (Figure 2B,C). The adhesion of platelets to PRCFC and PRPCR was specific as no adhesion was detected on HSA coated chambers even with residual fibrinogen and VWF present in perfused whole blood (Figure 2A–C).

FIGURE 2.

Platelet adhesion to PRCFC and PRPCR at low shear rate (300 s⁻¹). (A–C) Hirudinated whole blood was perfused into microfluidic chambers coated with PRCFC, PRPCR or 1% HSA for 5 min at 300 s⁻¹ (n = 7 blood donors). (A) Representative images of the microfluidic chambers after 5 min of blood perfusion. Scale bar = 30 μm. (B) Kinetics of platelet adhesion. (C) Numbers of adherent platelets were quantified at 5 min. *Symbols indicate the level of statistical significance by one‐way analysis of variance, Bonferroni post hoc test.

Previously, we showed that platelet adhesion to PRCFC is largely mediated by fibrinogen and VWF. ³³ To assess the respective contribution of these ligands to platelet adhesion on PRPCR, we repeated the experiments using whole blood treated with either abciximab, an αIIbβ3 integrin antagonist, or caplacizumab, a VWF inhibitor. Abciximab completely abolished platelet adhesion to both PRCFC and PRPCR (PRCFC − abciximab: 29,245 ± 2919 plts/mm² vs. PRCFC + abciximab: 830 ± 187 plts/mm²; PRPCR − abciximab: 18,612 ± 3149 plts/mm² vs. PRPCR + abciximab: 53 ± 15 plts/mm²; n = 4 blood donors), underscoring the central role of αIIbβ3‐fibrinogen interactions and supporting the observation that fibrinogen from PRCFC or PRPCR passively adsorbs onto the microfluidic chamber surface (Figure 3A,B). In comparison, caplacizumab caused only a slight, non‐significant reduction in platelet adhesion on PRPCR coatings (PRPCR without caplacizumab: 4885 ± 1055 plts/mm² vs. PRPCR + caplacizumab: 2493 ± 1371 plts/mm²; n = 4 blood donors). In contrast, caplacizumab had a larger effect on PRCFC coatings (PRCFC without caplacizumab: 15,324 ± 2595 plts/mm² vs. PRCFC + caplacizumab: 5166 ± 1496 plts/mm²; n = 4 blood donors), indicating the minimal contribution of VWF to platelet adhesion in PRPCR compared to PRCFC under these conditions (Figure 3C,D).

FIGURE 3.

Fibrinogen and not VWF are important for platelet adhesion to PRPCR at low shear rate (300 s⁻¹). (A, B) Microfluidic chambers were coated with PRCFC, PRPCR or HSA. 10 min before perfusion, blood was treated or not with abciximab (40 μg/mL). Hirudinated whole blood was perfused at 300 s⁻¹ for 5 min (n = 4 blood donors). *Symbols indicate the level of statistical significance by one‐way analysis of variance, Bonferroni post hoc test. (A) Numbers of adherent platelets on PRCFC or HSA were quantified at 5 min (B) Numbers of adherent platelets on PRPCR or HSA were quantified at 5 min. (C, D) Microfluidic chambers were coated with PRCFC, PRPCR or HSA. just before perfusion, blood was treated or not with caplacizumab (10 μg/mL). Hirudinated whole blood was perfused at 300 s⁻¹ for 5 min (n = 4 blood donors). *Symbols indicate the level of statistical significance by one‐way analysis of variance, Bonferroni post hoc test. (C) Numbers of adherent platelets on PRCFC or HSA were quantified at 5 min. (D) Numbers of adherent platelets on PRPCR or HSA were quantified at 5 min.

3.3. Contribution of VWF present in PRPCR to support platelet adhesion under high shear flow

To assess more specifically the contribution of VWF to platelet adhesion with PRCFC and PRPCR, we perfused reconstituted plasma‐free blood supplemented with either PRCFC or PRPCR, adjusted to a final fibrinogen concentration of 0.5 mg/mL, through microfluidic chambers coated with a VWF binding peptide (100 μg/mL) for 3 min at 1500 s⁻¹, to evaluate the adsorption and unfolding of circulating VWF. ³⁶ This wall shear rates was used in our study as it exceeds the threshold at which the GPIb‐IX‐V‐VWF interaction becomes crucial for platelet adhesion to collagen over 1000 s⁻¹. ³⁹ VWF in PRCFC demonstrated binding to the VWF binding peptide resulting in effective adhesion of platelets under an arteriole wall shear rate of 1500 s⁻¹ (PRCFC: 14,496 ± 2299 plts/mm²; n = 6 blood donors) (Figure 4A). In contrast, PRPCR lacking VWF and the negative control Tyrode's buffer did not support platelet adhesion (PRPCR: 228 ± 55 plts/mm²; n = 6 blood donors) (Figure 4B,C) indicating that the residual VWF in PRPCR was not functional.

FIGURE 4.

Platelet adhesion to VWF‐binding peptide at high wall shear rate (1500 s⁻¹). (A–C) Reconstituted blood (washed red blood cells + washed platelets + PRCFC, PRPCR or Tyrode's buffer) was perfused through microfluidic chambers coated with VWF‐binding peptide (100 μg/mL) for 3 min at 1500 s⁻¹ (n = 6 blood donors). (A) Representative images of the microfluidic chamber after 3 min of perfusion. Scale bar: 30 μm. (B) Kinetics of platelet adhesion. (C) Numbers of adherent platelets were quantitated at 3 min. *Symbols indicate the level of statistical significance by one‐way analysis of variance, Bonferroni post hoc test.

3.4. Functional properties of PRPCR to support platelet aggregation on collagen surfaces under shear flow

To simulate conditions in which endothelial cells are damaged with exposure of sub‐endothelial collagen, we examined the adhesion and aggregation of platelets on collagen‐coated microfluidic surfaces. Plasma‐free blood reconstituted with either PRCFC, PRPCR, or Tyrode's buffer was perfused at an elevated arterial wall shear rate (3000 s⁻¹) for 5 min to reproduce flow conditions found in wounds and in pathological conditions with vascular stenosis (Figure 5A–C). Plasma‐free blood reconstituted with PRPCR (fibrinogen 0.5 mg/mL and VWF <0.10 IU/mL) was unable to support platelet thrombus formation (PRPCR: 1139 ± 330 μm³; n = 6 blood donors), confirmed with the Tyrode's buffer control demonstrating a non‐significative difference between the two conditions (Figure 5B,C). In contrast, plasma‐free blood reconstituted with PRCFC, containing low levels of fibrinogen (0.5 mg/mL and VWF ~0.28 IU/mL), supported substantial platelet thrombus formation (PRCFC: 28556 ± 6576 μm³; n = 6 blood donors) (Figure 5A–C). These results indicate that PRPCR was unable to promote thrombus formation, due to markedly reduced functional VWF activity. In contrast, PRCFC, with retained levels of fibrinogen and VWF activity, supported platelet thrombus formation in the presence of hirudin inhibition of thrombin formation.

FIGURE 5.

Platelet aggregation on collagen surfaces at high wall shear rate (3000 s⁻¹). (A–C) Reconstituted blood (washed red blood cells + washed platelets + PRCFC, PRPCR or Tyrode's buffer) was perfused into microfluidic chambers coated with collagen HORM (200 μg/mL) for 5 min at 3000 s⁻¹ (n = 6 blood donors). (A) Representative confocal Z stack images of thrombi on collagen surface after 5 min of perfusion at 3000 s⁻¹. Scale bar: 50 μm. (B) Kinetics of thrombus formation. (C) Volume of thrombi was quantified at 5 min. *Symbols indicate the level of statistical significance by one‐way analysis of variance, Bonferroni post hoc test.

4. DISCUSSION

In this study, we evaluated the ability of PRPCR to support platelet functions under physiologically relevant shear conditions. PRPCR was markedly depleted in prothrombotic proteins VWF, fibrinogen, and FVIII associated with cryoprecipitate but retained a limited capacity to support platelet adhesion to fibrinogen at low shear rates while not supporting platelet aggregation on collagen at high shear rates. Importantly, PRPCR preserved thrombin‐generating capacity indicating a possibility to support the plasma phase of hemostasis. Moreover, PRPCR contained ADAMTS13 with substantial levels of intact immunoglobulins. Collectively, these findings support PRPCR as a pathogen‐reduced replacement fluid with features well suited for TPE, particularly in TTP.

McGuckin et al. reported that during the early recovery phase of TPE with rapidly rising platelet counts, anti‐platelet and anti‐coagulant therapy might be required to prevent thrombotic events. ²⁵ Therefore, in the current report we have extended the characterization of PRPCR using microfluidic assays to evaluate specifically the functional capacity of the retained fibrinogen and VWF on platelet adhesion and aggregation under clinically relevant wall shear rates. We selected PRCFC enriched for fibrinogen and VWF as a positive control adjusted to a fibrinogen concentration of 0.5 mg/mL in comparison to PRPCR because we could dilute PRCFC to minimal fibrinogen levels sufficient to conserve platelet aggregation with retention of sufficiently low levels of VWF to confirm microfluidic assay sensitivity to measure VWF function. Our study demonstrates that depletion of functional VWF in PRPCR did not substantially impact platelet adhesion on immobilized PRPCR at low wall shear rates (300 s⁻¹), which is explained by the ability of fibrinogen to support platelet adhesion. However, at high wall shear rates (3000 s⁻¹) requiring GPIb‐IX‐V complex‐VWF interaction, PRPCR demonstrated minimal platelet aggregation and thrombus formation because of the absence of functional VWF. This could be beneficial in patients requiring TPE to decrease the risk of platelet‐mediated thrombotic events in vascular access devices with elevated shear rates during rapidly rising or normal platelet counts. Importantly, PRPCR retains sufficient coagulation factor activity and anti‐thrombotic proteins to provide balanced hemostasis, and with sufficient levels of IgG, IgM, and IgA to avoid hypogammaglobulinemia during repeated TPE.

The other indications for TPE include a spectrum of malignant hematology disorders including monoclonal gammopathy hyper‐viscosity syndromes, non‐TTP microangiopathic syndromes, immune mediated hematologic disorders, solid organ transplantation desensitization and rejection, hematopoietic stem cell transplantation desensitization, neurologic disorders, and renal disorders. ⁶ The fluid replacement for the non‐TTP indications has generally been 5% albumin to avoid TTI risk, but extensive albumin replacement may require plasma infusion to correct coagulation and immunoglobulin deficits. Under these conditions, PRPCR may be beneficial for TPE to reduce TTI risk and thrombotic potential while retaining residual hemostatic capacity and measurable immunoglobulin content. IgG functionality was confirmed in the parent PR‐plasma from which PRPCR is derived, ⁴⁰ , ⁴¹ , ⁴² and CPP is a recognized starting material for plasma fractionation into immunoglobulins. ⁴³ Thus, we expect IgG function to be largely preserved in PRPCR; however, this remains to be demonstrated with direct functional assays on PRPCR.

A relevant consideration for therapeutic use of PRPCR is that the pathogen reduction process could potentially alter the quaternary structure of critical biological molecules. Prior publications have shown the retention of coagulation factor activity, including thrombin generation, in pathogen reduced plasma within therapeutic ranges. ³² , ⁴⁴ Proteomic analysis with quantitative two‐dimensional difference gel electrophoresis (2D‐DIGE) demonstrated modification of only four minor proteins not associated with coagulation function. In the current study, pathogen reduced cryoprecipitate fibrinogen complex (PRCFC) served as a positive control for PRPCR to characterize the depletion of fibrinogen and von Willebrand factor (VWF) in PRPCR. These studies demonstrate that fibrinogen and VWF in PRCFC supported the adhesion and aggregation of platelets under physiological conditions under low wall shear rates (300 s⁻¹) and under pathological conditions presenting high wall shear rates (3000 s⁻¹) suggesting that these complex molecules had not been substantially altered and were still functional.

A limitation of the study of PRPCR is that it has not been evaluated in clinical trials. However, there is extensive clinical experience with amotosalen‐UVA PR plasma, the source material for manufacture of PRPCR, for support of patients with congenital and acquired coagulopathies as well as TTP. In these studies, PR plasma demonstrated efficacy and safety sufficient for regulatory licensure with multiple years of safety experience. ¹⁷ , ⁴⁵ , ⁴⁶ , ⁴⁷ , ⁴⁸ Combined with this prior clinical experience and the experience with conventional CPP, our study supports the use of PRPCR in post‐marketing clinical studies of TTP and other TPE indications. However, the results obtained with the PRCPR in this model should be correlated with clinical outcomes in phase 4 studies with PRPCR in routine clinical use.

In conclusion, PRPCR is indicated to reduce the risk of TTI with provision of balanced hemostatic capacity, retained immunoglobulins, and reduced platelet thrombotic potential. Based on prior observations of clinical efficacy with non‐pathogen reduced CPP, we believe that PRPCR offers potential as an optimized replacement fluid for patients with TTP and for supplementation of patients undergoing TPE with 5% albumin for other indications where plasma transfusion may be required to correct coagulation and immunoglobulin deficits.

AUTHOR CONTRIBUTIONS

Florian Tupin, Pierre H. Mangin, and Beatrice Hechler designed the microfluidic assays. Clarisse Mouriaux performed the microfluidic assays. Kaja Kaastrup supervised and conducted the coagulation factor assays. Subra Yegneswaran supervised the manufacture of the blood components (PRPCR, PRCFC, and PR Plasma) used in these studies. Laurence Corash and Pierre H. Mangin contributed to the design of the studies. All of the authors participated in the writing and editing of the manuscript.

FUNDING INFORMATION

This study was supported by a contract from CERUS Corporation (Concord, CA), and funding from INSERM, EFS, and ARMESA (Association de Recherche et Développement en Médecine et Santé Publique).

CONFLICT OF INTEREST STATEMENT

Laurence Corash, Kaja Kaastrup, and Subra Yegneswaran are employees of Cerus Corporation and beneficial owners of Cerus equity. Pathogen Reduced Plasma, Cryoprecipitate Reduced is a product of Cerus Corporation marketed in the United States.

Supporting information

Data S1. Supporting Information.

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available from the corresponding author upon reasonable request.