US Military CPP Research and Development Programs

As an alternative to RT platelets, the US Office of Naval Research sponsored a CPP research program at the US Naval Blood Research Laboratory, Boston, MA, in the 1970s. Valeri and colleagues extensively characterized CPP in DMSO and evaluated in vitro and in vivo performance prior to advancing to clinical trials [32, 48, 50, 51]. A variety of parameters were measured including oxygen consumption, aggregation, release reactions, storage duration, ultrastructural alterations, and phagocytic function followed by evaluations of circulation and hemostatic effectiveness in canine, baboon, and other animal models [32, 48, 50, 51]. The Valeri method began with freezing single units of platelets prepared from WB in 5% DMSO at a freezing rate of 1 °C/min to −40 °C, followed by storage in the vapor phase of liquid nitrogen [52]. Frozen platelets were prepared for transfusion by thawing in a 37 °C water bath, washing, and resuspending in plasma. Valeri’s group continued to develop and refine their methods ending with freezing of apheresis collected platelets in 6% DMSO that were frozen and stored at −80 °C. The units were thawed, washed, and resuspended in anticoagulant citrate dextrose (ACD) solution plasma prior to transfusion.

The culmination of this development program was a clinical trial in patients undergoing cardiopulmonary bypass surgery who were randomized to receive either CPP or RT platelets in the perioperative period. The trial was successful in demonstrating that CPP established hemostasis as effectively as control platelets and non-inferiority metric was met [53]. The results were published in 1999, but the cumulative preclinical and clinical data on the Valeri method for platelet freezing were not sufficient to obtain FDA approval. Valeri and colleagues continued development including the pivotal invention of the no-wash method , enabled by removal of the DMSO/plasma supernatant prior to freezing [54].

International CPP Research and Development Programs

The Netherlands Armed Forces (NLAF) also experienced considerable challenges in supplying platelets to austere environments. Having learned from the Valeri laboratory, the NLAF chose to augment standard blood products with CPP and other frozen blood products rather than rely on a walking blood bank for fresh WB and apheresis platelet products. The NLAF began using frozen 6% DMSO platelets reconstituted in thawed plasma with thawed deglycerolized red cells and thawed fresh frozen plasma (FFP) in their deployed medical treatment facilities (MTFs) in 2001 [55]. They published their experience in 2016 after reviewing 4 years of hemovigilance and combat casualty outcomes data collected in Afghanistan beginning in 2006. Results were excellent, with only 1 observed mild transfusion reaction in 3060 transfused products. Twenty-four-hour and hospital stay mortality (14% compared to 44% in historical controls, p = 0.005) decreased significantly in massive transfusion patients. The authors concluded that the use of frozen products, including CPP, in massive transfusion is safe and effective [55].

Rapid dissemination of these findings was likely facilitated during the recent conflicts in Iraq and Afghanistan, because exchange of medical information was common among the North Atlantic Treaty Organization (NATO) coalition forces [56]. In Australia, the University of Queensland, the Australian Red Cross (ARC), the Australian Defense Force (ADF), and others are collaborating to expedite a CPP development program [57–59]. In addition to potential military applications, providing a platelet product with storage times of years, rather than days, could provide hemostatic platelets in smaller rural hospitals that cannot sustain an RT platelet concentrate inventory. One-third of Australia’s population lives in remote areas where providing emergency services and blood products is challenging [60]. The recognition that longer-stored, potentially more hemostatic platelet-derived products are needed for both civilian and military centers mirrors similar viewpoints recently expressed by physicians in the USA and Europe [42, 61].

The ARC has studied CPP using various preparations in platelet starting material. The major differences included the initial storage solution (70% platelet additive solution, or PAS, versus 100% plasma), the final reconstitution fluid (70% PAS versus 100% plasma), and method of collection (apheresis versus buffy coat preparation) [62–66]. ARC reported differences in factor activity, protein content, microparticles, and other biological response modifiers in these CPP compared to standard RT platelets [64]. They noted that fibrinogen was higher in CPP, whereas factors V and VIII activity were decreased. The clinical significance of these findings regarding in vivo CPP function and outcomes was difficult to estimate given that these platelets are primed for activation and in vitro studies are difficult to correlate to clinical findings. The ARC investigators found that thrombin generation and clotting activity were faster, while clot strength was similar to fresh [64–66]. As reported by others, aggregation was attenuated compared to Day 2, but comparison was not made to aggregation after Day 3, when RT platelets typically respond poorly to agonists.

Reade and colleagues recognized the limitations of currently available clinical data for CPP. The University of Queensland, the ARC , and the ADF collaborated on a randomized, controlled multicenter clinical trial in a surgical bleeding population, Cryopreserved Platelets Versus Liquid Platelets (CLIP) Trial (ACTRN12612001261808) [57]. The investigators hypothesize that CPP will be at least as effective and safe as conventional RT platelets in the management of active bleeding related to surgery. The primary endpoints are protocol feasibility in this setting and safety and acceptability [57]. Clinical efficacy as measured by 28-day mortality, blood loss, transfusion requirement, and thromboembolism will be assessed as secondary endpoints. Results are not yet available.

The US Army has sponsored a development program for CPP. Based on the Valeri no-wash method, a Current Good Manufacturing Practice (cGMP) manufacturing process has been developed [67]. Briefly, gamma-irradiated (25Gy) apheresis platelets in plasma from either the Trima apheresis system (Terumo BCT, Inc., Lakewood, CO, USA) or Amicus apheresis system (Fresenius-Kabi, Lake Zurich, IL, USA) are brought to 5.6 to 6.7% effective DMSO concentration by adding 27% sterile DMSO/saline concentrating by centrifugation at 1250× g for 10 min, removal of supernatant to a final volume of 20–35 mL in a freezing bag (Cryostore 500, OriGen Biomedical, Austin, TX, USA), placed into a polyvinyl chloride (PVC) overwrap bag, and stored in a cardboard freezing box. The box is placed flat on the floor of a chest-type mechanical 80 °C freezer. CPP are stable for up to 24 months at ≤ −65 °C. To prepare for transfusion, CPP is thawed in a 37 °C water bath for 8 min. After a 30-min rest period, 0.9% NaCl at 20–24 °C is added slowly in aliquots of 10 mL and 15 mL while gently mixing. The resuspended CPP contains approximately 2.0 × 10¹¹ to 3.6 × 10¹¹ irradiated platelets in a volume of ≥45 mL to ≤60 mL and approximately 1250–2530 mg residual DMSO. Thawed diluted CPP units are stable at RT for up to 4 h.

CPP In Vitro Phenotype

The phenotype of thawed CPP is distinct from RT 5-day liquid-stored platelets (LSP). CPP presented with increased microparticles content, phosphatidylserine exposure, and thrombin generation capabilities (thrombin peak, Table 9.1), suggestive of a more activated cell state. The more activated state of CPP is apparent in transmission electron micrographs (Fig. 9.2). Comparing fresh platelets (Fig. 9.2a) to CPP (Fig. 9.2b), while the ultrastructure of microtubules and open canalicular system is maintained in many cells, the CPP (Fig. 9.2b) cell population has a larger representation of active platelets. Approximately 30–50% of 4000× fields appear as platelets, with more pseudopodia observable, occasional platelets with granule contents constricted to the center of the cell, and some platelets observed with few, if any, granules (Fig. 9.2b). No aggregates, per se, were observed (Fig. 9.2b). Confirming the increased microparticle contents of thawed CPP identified with flow cytometry (Table 9.1), the 2000× g CPP supernatant prepared with the 16,000× g centrifugation shows many spherical particles 500 nm and smaller (Fig. 9.2c).

Table 9.1

In vitro phenotype for 5-day LSP and CPP

	5-day LSP	CPP
Platelet concentration (×103/μl)	1521 ± 144	6295 ± 816
Microparticles (% of platelet events)	6 ± 3	42 ± 9
Phosphatidylserine exposure (% of platelet events)	7 ± 4	73 ± 4
Aggregation (ADP + EPI) (%)	75 ± 24	20 ± 13
Aggregation (collagen) (%)	83 ± 14	16 ± 12
TGA peak thrombin (nM) (SD)	72.3 (10.3)	159.6 (25.7)

From Cid et al. [68], with permission of John Wiley and Sons

For aggregation assay final concentrations: ADP adenosine diphosphate at 10 μM, EPI epinephrine at 5 μM; collagen at 10 μg/ml and platelet concentrations at 300 × 10³ platelets/μl. For TGA thrombin generation assay with final concentration of 66 × 10³ platelets/μl, 1pM tissue factor in pooled fresh frozen plasma

LPS liquid-stored platelet, SD standard deviation

Table 9.2

Transfused platelet recovery and survival

	Platelet recovery (%)	Platelet survival (d)
Fresh	63 ± 9	8.6 ± 1.1
CPP	33 ± 10	7.5 ± 1.2
P	<.0001	<.0001
% of fresh	52 ± 12	89 ± 15

From Slichter et al. [70], with permission of Elsevier

Data reported as mean ± standard deviation, n = 32

Interestingly, CPP response to exogenous agonist compared to 5-day LSP in an aggregation assay is decreased (Table 9.1). Examining the clotting phenotype of CPP under shear flow over rabbit aorta in a flow chamber (with CPP restored platelet-depleted WB) revealed CPP maintained half of the platelet deposition capability of WB restored with 5-day LSP under medium shear (Fig. 9.3, CPP at 10.7 ± 3.1 and 5-day LSP at 22.9 ± 4.1 percent platelet coverage) [68]. Fibrin coverage is equivalent between both cell types (Fig. 9.3, 21–25%); however, CPP supported an almost threefold higher prothrombin cleavage which is maintained over 10 min of shear flow (Fig. 9.3, CPP at 606 ± 216 compared to LSP 221 ± 67) [68].

CPP Recovery and Survival

Autologous recovery and survival of CPP in healthy subjects have been evaluated compared to fresh autologous platelets transfused simultaneously following the method recommended by the Biomedical Excellence for Safer Transfusions (BEST) Collaborative, with some modifications for CPP [67, 69, 70]. As expected with an activated platelet product, CPP recovery was 52% of fresh platelets (Table 9.2); however, they circulated for 7.5 days or 89% of fresh platelets [70]. This is consistent with good clinical outcomes previously reported when similar CPP products were used for hypoproliferative thrombocytopenia as discussed above. CPP recoveries are lower and could result in the need for increased transfusions, but it is also possible that CPP may allow physicians to manage patients with a lower platelet count due to the improved hemostatic potential of this product. Further studies are needed to evaluate CPP safety and clinical utility, and these are currently in progress.

CPP Phase 1 Dose Escalation

Advancing CPP into clinical trials has been challenging. In a Phase 1 clinical trial (Safety Study of Dimethyl Sulfoxide Cryopreserved Platelets; ClinicalTrials.​gov Identifier: NCT02078284) [71], patients with a WHO bleeding score ≥2 received from 0.5 to 3 units of CPP (n = 24) or 1 unit of LSP (n = 4). There were no related thrombotic or other serious adverse events and five mild transfusion-related adverse events. Among the CPP recipients, 14/24 (58%) had improved bleeding scores, including 3/7 (43%) patients who had intracerebral bleeding. CPP post-transfusion platelet increments were significantly less than LSP (Fig. 9.4); however, days to the next transfusion for CPP or LSP were the same [71]. A Phase 2 trial in cardiac surgery patients undergoing coronary artery bypass grafting (CABG) is planned for the near future (Fig. 9.1).