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Frozen red cells for military and civil purposesRelevance, experiences and developmentsLelkens, C.C.M.

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Frozen Red Cells for M

ilitary and Civil Purposes C

.C.M

. LelkensC.C.M. Lelkens

Frozen Red Cells for Military and Civil Purposes

Relevance, Experiences and Developments

Twee erythrocyten uit Leiden

Besloten de warmte te mijden.

Naar de MBB,

Zo spraken de twee,

De diepvries zal ons gaan verblijden.

Colofon

ISBN (book): 978-90-826959-0-8

ISBN (digital document): 978-90-826959-1-5

Author: Charles Chrétien Marie Lelkens

Cover design: Evelien Jagtman, Maastricht

Print: Drukkerij Mostert en Van Onderen, Leiden

Copyright © 2017 Charles Lelkens

Alle rechten voorbehouden. Niets uit deze uitgave mag worden verveelvoudigd, opgeslagen in een geautomatiseerd gegevensbestand of openbaar gemaakt worden in enige vorm of op enige wijze, hetzij elektronisch, mechanisch of door fotokopieën, opname, of op enige andere manier, zonder voorafgaande schriftelijke toestemming van de auteur.

All rights reserved. No part of this publication may be reproduced, stored in retrieval systems, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the author.

Deze uitgave kwam tot stand dankzij een genereus gebaar van de schrijver.

Frozen Red Cells for Military and Civil Purposes

Relevance, Experiences and Developments

ACADEMISCH PROEFSCHRIFT

ter verkrijging van de graad van doctor aan de Universiteit van Amsterdamop gezag van de Rector Magnificus

prof. dr. ir. K.I.J. Maex

ten overstaan van een door het College voor Promoties ingestelde commissie,in het openbaar te verdedigen in de Aula der Universiteit

op vrijdag 30 juni 2017, te 11.00 uur

door

Charles Chrétien Marie Lelkens

geboren te Leiden

Promotiecommissie

Promotor: Prof. dr. A. J. Verhoeven AMC-UvACopromotores: Dr. J.W.M. Lagerberg Sanquin Research Dr. D. de Korte Sanquin Research

Overige leden: Prof. dr. E. van der Schoot AMC-UvA Prof. dr.ir. C. Ince AMC-UvA Prof. dr. T.M. van Gulik AMC-UvA Prof. dr. J.R. Hess University of Washington Dr. G.J.C.G.M. Bosman Radboudumc Dr. H. Woelders Wageningen University & Research

Faculteit der Geneeskunde

Aan mijn ouders, in liefdevolle en dankbare herinnering

Aan Jacqueline, Marie-Christine en Jean-Louis

“In God we trust, from others we need data”

Carlo Robert Valeri, MD (1932)Director Naval Blood Research Laboratory,

Plymouth, MA, USA

Contents

Chapter 1 Introduction

Chapter 2 Stability after thawing of RBCs frozen with the high- and low-glycerol method. (Transfusion. 2003; 43(2):157-64)

Chapter 3 Experiences with frozen blood products in the Netherlands military. (Transfus Apher Sci. 2006; 34(3):289-98)

Chapter 4 Australian experience with frozen blood products on military operations. (Med J Aust. 2010; 192(4):2035)

Chapter 5 Prolonged postthaw shelf life of red cells frozen without prefreeze removal of excess glycerol. (Vox Sang. 2015; 108(3):219-25)

Chapter 6 The effect of prefreeze rejuvenation on postthaw storage of red cells in AS-3 and SAGM. (Accepted by Transfusion)

Chapter 7 Advances in military, field, and austere transfusion medicine in the last decade. (Transfus Apher Sci. 2013; 49(3):380-6)

Chapter 8 General discussion

Chapter 9 Summary

Chapter 10 Samenvatting

Appendices Acknowledgments / Dankwoord Curriculum vitae auctoris Resume

11

25

45

67

79

95

119

137

153

159

165169171

Chapter

Introduction

1

“Mijn” schip: standaardfregat Hr.Ms. Banckert (1980-1981)bron: Nederlands Instituut voor Militaire Historie, beeldbank.

Introduction

13

Introduction

Large-scale military conflicts (or “wars”) have undeniably played an important role in the progress of medical care with, for instance, the ligature, the plaster cast and the wheel stretcher as obvious examples. Civilian medical practice today still benefits from these war-driven innovations, introduced by military doctors with battlefield experience. The same holds true for several developments in blood transfusion, with the use of frozen (or cryopreserved) red blood cells as most outstanding example. For this reason, the history of the development of cryopreserved red blood cells for transfusion will be described in this chapter according to the time scale determined by major military conflicts.

The US Civil WarThe first case reports of a successful military blood transfusion concerned two wounded soldiers in the US Civil War (1861-1865). In 1864, two Union Army surgeons, assigned to two different hospitals, wide apart, decided to transfuse two soldiers with human blood from healthy individuals after a leg amputation.1,2 Both soldiers survived.

In the 1890’s the potential of citrate as an anticoagulant was discovered, but not put into clinical practice until its independent rediscovery by three researchers in three different countries in 1914 and 1915.3,4 This paved the way for a safe preservation of blood for future use.

The First World War Although in April 1915 the first recorded transfusion of World War I (and of the 20th century) was administered,5 it was not before 1916 that Canadian Captain Lawrence Bruce Robertson published the first article about his experiences with transfusing (uncrossmatched) blood via syringe and cannula in war time circumstances.6 Despite the fact that another Canadian, Major Edward Archibald, decided to introduce the use of sodium citrate in 1915,7 that allowed him to transfuse blood when needed,8 Robertson preferred saline to flush syringe and cannula periodically. Like most surgeons at that time, he was very cautious in adding substances to blood. Until then, most transfusions were performed directly via end-to-end anastomosis of artery and vein of donor and recipient respectively.9 This technique proved to be totally unsuitable for emergency purposes, because it not only required two skilled surgeons and nursing staff,

Chapter 1

14

Introduction

15

but also took precious time and the proximity of an available donor in the operating room. In addition, there was always the danger of overdonation or undertransfusion.3,4,10-12

In the first year of World War I, the US Army ordered further exploration of this field, leading to experiments by Rous and Turner, who in 1916 published their findings, indicating that a mixture of 5.4% glucose and 3.8% sodium citrate could protect human red blood cells from hemolysis for four weeks.13 In April 1917, the United States entered the war and during the battle of Cambrai at the front in France in November 1917, a US military doctor, Captain Oswald H. Robertson, successfully applied the Rous-Turner solution, achieving up to 26 days of storage of whole blood.4,14 He used a self-designed icebox,14,15 containing glass bottled whole blood, thus becoming the world’s first blood banker.14 Although there are no exact figures of how many transfusions were performed during World War I, several tens of thousands in British and Canadian hospitals in 1918 appears to be a fair estimate.12

The Spanish Civil WarAlmost two decades later, during the Spanish Civil War (1936-39), for the first time in history, the concept was initiated to collect and store (civilian) citrated and typed blood in a (civilian) blood bank16,17 and take it from there to the patient as close to the front as possible.18-20 The two most famous civilian blood banks were set up in Barcelona and Madrid. The transfusion service in Barcelona alone collected, processed, tested and distributed some 9,000 liters of whole blood between January 1936 and August 1939, using only group O from over 28,000 donors,16,17,21 resulting in 27,000 transfusions.

Between the two World Wars, in 1928, dr. Shamov in Ukraine suggested human cadaver blood as a safe source for transfusion. His idea was put into practice in 1930 by a Russian colleague, dr. Sergei Yudin, who in 1937 reported on having performed 1000 successful transfusions, without using citrate but relying on post mortem fibrinolysis instead.22,23 During a visit to Spain in 1934, Yudin pointed out the convenience of his idea, but it was probably never put into practice in the course of the war, at least not at the Barcelona institute.20,24 Although logical in concept, moral, ethical, but also practical consequences,22,25,26 prevented this idea from gaining solid ground in day-to-day clinical practice outside the Soviet Union, even in war time conditions, but it stimulated further development of blood conservation and blood banks.27

Between 1938 and 1940, the Russians used anticoagulated donor blood in military operations on the Kuril Islands (1938) and in Finland (1939-1940), prepared at blood transfusion institutes in Leningrad and Moscow. So when Russia entered World War II, the blood supply system was in place and, in the end, able to provide the armed forces with a total of 1.7 million liters of blood along the front lines during the war.28

The Second World War and the Korean WarThe experiences in Spain and Russia in the late 1930s finally led to the deployment of blood transfusion services in World War II, capable of large-scale collections and storage of whole blood. In 1940, the work of Cohn enabled the plasma fractionation of whole blood and when the Japanese attacked Pearl Harbor, albumin was available and immediately shipped to Hawaii to treat 87 victims, mainly burn patients, of which some showed dramatic improvement.10 Although some doctors, based on their World War I experiences, had learned that severe hemorrhage led to shock and therefore had expressed their opinion that oxygen carrying capacity was the primary need, the general belief (at least in the US) was that with the availability of plasma (liquid or freeze-dried) this alone was sufficient to compensate for the blood loss. Other than that, blood transfusion was considered too difficult and dangerous and its supply was logistically much more complicated.21 Emphasis was therefore laid on the use of plasma by the US forces at the start of World War II. It was only in 1943, because of reports from the North-African theater,29 as well as the situation in Europe and in the Pacific in 1944,12 that attention shifted back again to whole blood.21 This persisted through the Korean War (1950-1953), during which almost unlimited amounts of whole blood were available and used to the maximum.29 Adequate resuscitation, among which the availability of blood, played an important role in reducing the numbers of wounded servicemen dying after reaching the hospital. During World War I, around 10% of those arriving at a hospital died, in World War II this number had decreased to 4.5% and finally in the Korean War to 2.6%.21

The Vietnam WarSoon after their direct military involvement in the Vietnam War (1946-1975) the US started to provide whole blood to Saigon (South Vietnam) in 1965. In that same year and the year thereafter, packed RBC and fresh frozen plasma (FFP) became largely available. Until 1971 approximately 1.3 million red

Chapter 1

16

Introduction

17

cell units were sent, of which, due to outdating, less than half (600,000, i.e. only 46%!) were administered in US hospitals. Partly because of this, the US Department of Defense (DoD) sponsored clinical research towards extension of the storage period of RBC, without yielding a licensed product though.12 Despite developments over the past decades since World War II, current preservation solutions are still not approved beyond 42 days.30 Substantial extension of (hypothermic) storage periods is possible by using cryopreservation techniques, i.e. storage below 0°C. Currently, frozen red cells can be stored for at least 10 years, according to European and US guidelines.30,31

Development of cryopreservation techniques Commissioned by the US Navy in 1956, the Blood Research Laboratory (in 1965 renamed as the Naval Blood Research Laboratory) started to pursue another way to prolong the storage period of RBC,32 based on the findings of dr. Audrey Smith.33 She had observed that high concentrations of glycerol (45-50% w/v) could serve as a cryoprotectant for red cells, thus enabling their preservation by freezing and storing at temperatures below -65°C. In the early sixties, several investigators34-36 found that lower concentrations of glycerol (17-20% w/v), were able to act as cryoprotectant, but only if the rate of freezing was accelerated by immersion in liquid nitrogen, followed by storage below -150°C in (the vapor phase of) liquid nitrogen. These two methods, referred to as the high- (HGM) and low-glycerol method (LGM) respectively, are still in use to freeze red cells.

The low rate of freezing and the possibility to store at relatively high temperatures (below -65°C), makes the HGM the more practical cryopreservation method: cells can be frozen and stored in mechanical freezers and transported on “dry ice”. The obligatory use of liquid nitrogen precludes the LGM from being used in an operational, military setting, leaving the HGM as the only method to extend shelf lives of blood products for military purposes.

Cryoprotection by glycerol and its pretransfusion removalGlycerol is most effective in protecting those cells into which it permeates fairly rapidly. Human erythrocytes have a high permeability for glycerol, unlike erythrocytes of pigs, dogs and cats for instance.37 Uptake of glycerol occurs both by active (or facilitated) and passive diffusion.38-40 The former is a metabolic process and as such temperature dependent. Passive diffusion is essentially independent of temperature.38 Glycerol and other intracellular cryoprotectants

form very firm hydrogen bonds with intracellular water, preventing this captured water turning into ice. With the available water for ice crystallization minimized, glycerol suppresses the rise in NaCl concentration, thereby avoiding extreme hypertonicity.41-43 Glycerol is thus effective in preventing cellular lysis due to osmotic damage during freezing and thawing.38

Regardless of the glycerol concentration used, postthaw washing of cryopreserved RBC is necessary to lower the concentration of glycerol to less than 1% (w/v) before transfusion.44 On contact with (isotonic) plasma, the glycerolized cells would otherwise hemolyze, because water enters the red cell faster than glycerol is able to move out.45 This would cause swelling and eventually hemolysis if no precautions were taken. Different methods have been developed and used to achieve acceptable concentrations of glycerol in thawed red cell preparations, like dilution, dialysis and serial or continuous washing.

In 1954 the Cohn fractionator, originally designed to extract protein fractions from blood plasma, was modified to add and remove glycerol.46 The device used a continuous centrifugation process with a gradient hypertonic electrolyte wash of glycerol, sodium lactate and saline. Although successful, it proved to be a complicated, time consuming and impractical technique for widespread clinical use.47

In 1963 Huggins introduced a simpler method (without centrifugation) to remove glycerol, the so-called “reversible agglomeration”, to distinguish the phenomenon from agglutination and aggregation.48 The procedure removes the glycerol by adding large volumes of non-electrolyte solutions containing glucose and fructose, while stirring. As a result, the red cell environment is low in ionic strength and the cells start to clump spontaneously and settle as soon as stirring is stopped. The supernatant is then decanted and disaggregation of the retained red cell mass is then achieved by the addition of electrolyte solutions, like isotonic saline.41 Prior to transfusion, the red cells need to be concentrated to obtain a final hematocrit of 90%.49

Despite extensive and favorable (clinical) experience with this technique, gained at the US Naval hospital in Chelsea (MA)50-52 and, later, during the Vietnam War,53,54 the main source in the Vietnam War remained red cells prepared without freezing. However, between 1966 and 1969, the US Navy in Da Nang used more than 2,000 units of frozen red cells (O pos and O neg) with satisfactory results.55 The major logistic problems with the Huggins-method proved to be the large volume of washing solution and deglycerolization time.56

Chapter 1

18

Introduction

19

Moreover, after storage at -80°C for 18 months or longer, the RBC recovery values were only around 60%. Altogether, these drawbacks outweighed the shorter shelf life of the abundantly available fresh blood and the frozen blood banks were withdrawn from Vietnam in 1972. The unused frozen units were transported back to the US in styrofoam containers filled with dry ice and used for further research purposes.57

In 1967, the Arthur D. Little Company introduced the (reusable) stainless steel bowl,58 later followed by the (disposable) polycarbonate version in several generations of Haemonetics blood processors.59 Both bowls were used for continuous-flow washing and have been extensively tested by Valeri and associates.60-65 The M115 cell processor (Haemonetics, 1976), with a polycarbonate bowl, had an integrally attached shaker, to ensure adequate mixing of the various washing solutions. It was actually deployed and used during the first Gulf War (1990-1991). Deglycerolization took only about 35 minutes and 2L of washing fluids,66 in contrast to the Huggins method, taking 50 minutes and 6.8 L.55 It was still an open system, though, so the thawed red cells had to be administered within 24 hr after thawing and washing, in conjunction with storage at 2-6°C.

The practical use of frozen red cells during conflicts in the past two decadesIn 1990, during the build-up to the first Gulf War, some 7,000 frozen units were shipped to the area of conflict, of which only 265 were deglycerolized, without any transfusion. In sharp contrast, while preparing for this war during Desert Shield, some 82,000 liquid red cell units were shipped to the area of conflict, of which around 250 units were used for 250 US casualties and another 750 units for Iraqi prisoners of war (POW) and civilians. Ultimately, around 67,000 liquid red cell units, 80 percent of what was sent, were not used and had to be discarded.12 This outcome closely resembles the fate of the 3262 units of liquid red cells, provided to support the British armed forces in the Falklands War in 1982. Only 605 were used, giving a usage rate of 18.5 %.67

In 1991, the Netherlands military took part in the first Gulf War, mainly by deploying a navy task force, which was supplied with regular shipments of fresh red cells (2-6°C). The same array of products was used during the 1992-1993 United Nations-mission in Cambodia (UNTAC).

The experiences obtained from these two missions showed that worldwide deployments create serious logistic challenges, including those regarding blood

supply. Considerable distances need to be bridged between the Netherlands and the area of operations. Other than the fact that this takes precious time, the quantities and points in time where blood products are needed, cannot be predicted in wartime conditions. The relatively short shelf lives of particularly platelets and red cells would require frequent shipments. However, the points in time where and how much of the necessary blood products are needed would still remain unpredictable. So, even if at all times all necessary blood products would be made available, using the standard storage techniques, the operational needs would only be covered against considerable costs. At the same time, there would still be the danger of having to discard substantial numbers of units, due to outdating.12,67 This adds an ethical aspect to the discussion, because all blood products have been provided by volunteer, non-remunerated donors, aiming at helping those in need of transfusions. The only storage method, currently available, that guarantees the highest degree of self-sufficiency possible for the deployed military units, with the lowest outdating rates, is the application of cryopreservation. The first steps of the Netherlands military towards a blood supply system based on frozen blood products were taken during the UN- and NATO-missions in Bosnia (1992-1995 and 1995-2004 respectively), during which frozen red cells (using the HGM according to Valeri), FFP and eventually, by the end of 2001, frozen platelets in dimethyl sulfoxide (DMSO), all stored at - 80°C, became available to treat hemorrhaging war casualties. Deglycerolization of the thawed red cells was still performed with the (Haemonetics) M115, therefore allowing only for 24 hr of postthaw storage. After the introduction of the fully closed, automated cell processor ACP 215 in 1998, it became possible to extend the postthaw shelf life of thawed red cells to 14 days, also because of the application of AS-3 (Nutricel®) as the final storage solution.68 The Netherlands, taking part in missions in Iraq (2003-2005) and Afghanistan (2002-2014), gained extensive experience with shipping, storing, preparing and administering (previously) frozen blood products in war time conditions, achieving a safe and effective blood supply under wartime conditions, with a minimum wastage rate due to outdating and a minimal burden to the logistic system.69,70

In the next chapters the developments in the Military Blood Bank and research efforts to improve the practicality of the use of frozen red cells in battlefield (and civil) conditions will be discussed.

Chapter 1

20

Introduction

21

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55. Valeri C.R.and Gina Ragno Giorgio. The US Navy’s experience with resuscitation of wounded servicemen in Vietnam using frozen washed red blood cells from 1966 to 1974: developments from this experience. In Forty-five years of Research at the NBRL, Boston, Massachusetts. first ed. 2013. p. 208-9.

56. Valeri C.R.and Gina Ragno Giorgio. The US Navy’s experience with resuscitation of wounded servicemen in Vietnam using frozen washed red blood cells from 1966 to 1974: developments from this experience. In Forty-five years of Research at the NBRL, Boston, Massachusetts. first ed. 2013. p. 204.

57. Valeri CR. Factors which affect the therapeutic effectiveness of red cells freeze-preserved with glycerol. In Blood Banking and the Use of Frozen Blood Products. CRC Press; 1976. p. 54.

58. Tullis JL, Tinch RJ, Gibson JG, Baudanza P. A simplified centrifuge for the separation and processing of blood cells. Transfusion 1967;7(3):232-42.

59. Tullis JL, Gibson JG, Tinch RJ, Hinman J, Baudanza P, DiForte S, Smith T, Breed AT. Disposable plastic centrifuge bowls for separation of red blood cells and plasma in the processing of frozen blood. Transfusion 1971;11(6):358-67.

60. Runck AH, Valeri CR, Sampson WT. Comparison of the effects of ionic and non-ionic solutions on the volume and intracellular potassium of frozen and non-frozen human red cells. Transfusion 1968;8(1):9-18.

61. Valeri CR, Runck AH. Long term frozen storage of human red blood cells: studies in vivo and in vitro of autologous red blood cells preserved up to six years with high concentrations of glycerol. Transfusion 1969;9(1):5-14.

62. Runck AH, Valeri CR. Recovery of glycerolized red blood cells frozen in liquid nitrogen. Transfusion 1969;9(6):297-305.

63. Crowley JP, Valeri CR. The purification of red cells for transfusion by freeze preservation and washing. I. The mechanism of leukocyte removal from washed, freeze-preserved red cells. Transfusion 1974;14(3):188-95.

64. Crowley JP, Valeri CR. The purification of red cells of transfusion by freeze preservation and washing. II. The residual leukocytes, platelets, and plasma in washed,freeze-preserved red cells. Transfusion 1974;14(3):196-202.

65. Crowley JP, Valeri CR. The purification of red cells for transfusion by freeze-preservation and washing. III. Leukocyte removal and red cell recovery after red cell freeze-preservation by the high or low glycerol concentration method. Transfusion 1974;14(6):590-4.

66. Valeri CR, Valeri DA, Anastasi J, Vecchione JJ, Dennis RC, Emerson CP. Freezing in the primary polyvinylchloride plastic collection bag: a new system for preparing and freezing nonrejuvenated and rejuvenated red blood cells. Transfusion 1981;21(2):138-49.

67. Marsh AR. A short but distant war - the Falklands campaign. J.R.Soc Med 1983;76(11):972-82.

68. Valeri CR, Ragno G, Pivacek LE, Srey R, Hess JR, Lippert LE, Mettille F, Fahie R, O’Neill EM, Szymanski IO. A multicenter study of in vitro and in vivo values in human RBCs frozen with 40-percent (wt/vol) glycerol and stored after deglycerolization for 15 days at 4 degrees C in AS-3: assessment of RBC processing in the ACP 215. Transfusion 2001;41(7):933-9.

69. Lelkens CCM, Koning JG, de Kort B, Floot IBG, Noorman F. Experiences with frozen blood products in the Netherlands military. Transfus.Apher.Sci. 2006;34(3):289-98.

70. Noorman F, van Dongen TTCF, Plat MC, Badloe JF, Hess JR, Hoencamp R. Transfusion: -80 degrees C Frozen Blood Products Are Safe and Effective in Military Casualty Care. PLoS.One. 2016;11(12):e0168401

Charles C.M. Lelkens1, Femke Noorman1, Jack G. Koning1,Rosa Truijens-de Lange2, Perry S. Stekkinger2, Joa C. Bakker2,

Johan W.M. Lagerberg2, Anneke Brand3 and Arthur J. Verhoeven2

1Military Blood Bank, Leiden, 2Department of Transfusion Technology, Sanquin Research at CLB, Amsterdam,

3Blood Bank Sanquin South-West, Leiden

Transfusion. 2003;43(2):157-64

Chapter

Stability after thawing of RBCs frozen with the high- and low-glycerol method

2

Chapter 2 Stability after thawing of RBCs frozen with the high- and low-glycerol method

26 27

Abstract

Background: RBCs can be frozen with either the high-glycerol method (HGM) or the low-glycerol method (LGM). To date, the use of frozen RBCs is hampered by a 24 hour outdating period after thawing. A closed washing system (ACP 215) may solve this problem.

Study design and methods: We compared the effects of high- (40%) and low-glycerol (19%) concentration, with and without freezing (at -80°C for HGM, -196°C for LGM) on the in vitro quality of RBCs after deglycerolization with the closed washing system and during storage at 4°C in SAGM after thawing.

Results: Glycerol treatment by itself induced hemolysis during processing, which was more pronounced in HGM cells. The freeze-thaw-wash process decreased the stability of RBCs, particularly in LGM cells during storage after thawing. In contrast to LGM cells, in HGM cells no additional effect of freeze or thaw on stability of washed cells was seen during the first week of storage after thawing. Changes in osmotic resistance and cellular metabolism could not explain the observed differences in RBC stability.

Conclusion: The closed washing system is able to process both high- and low-glycerol-treated RBCs. Stability after washing during cold storage in SAGM, as measured by hemolysis, is better for HGM cells as compared to LGM cells.

Introduction

Freezing RBCs with glycerol as a cryoprotectant dates back to 19501 after an accidental discovery the previous year.2 In the 1960s, it was found that accelerating the rate of freezing could significantly reduce the required concentration of glycerol to 19-20%.3-5 After several modifications6-9 the two methods, referred to as the high- (HGM) and low-glycerol methods (LGM) respectively, are still in use for freezing units of RBCs. HGM allows storage at -80°C, whereas LGM requires liquid nitrogen (-196°C). Some laboratories prefer LGM over HGM because of a shorter processing time and a more or less indefinite shelf life at temperatures below -130°C.10,11 Recently, however, Valeri et al.12 have shown that cells frozen with HGM and stored at - 80°C can be stored for up to at least 37 years with an acceptable recovery after thawing and washing. Despite their undeniably higher costs,13-16 frozen RBCs have several advantages that are, paradoxically, mainly related to the necessary washing procedure, eliminating cell debris, WBCs, cytokines, and free Hb.17-19 Stockpiling a frozen inventory of RBCs can be done in the case of rare blood groups and for military deployments, which characterized by logistical problems and unpredictable needs of blood components.20 However, besides cost, the actual use of frozen RBCs is hampered mainly by processing time and a 24 hour outdating period due to potential bacterial contamination if thawing and washing is performed in a non-closed system.21,22 Recently, a fully automated and functionally closed washing system for RBCs has been introduced (ACP 215, Haemonetics, Braintree, MA), which may contribute to a solution of these problems.

Although a closed washing system has been developed for processing RBCs according to HGM,23-25 we investigated its application for LGM as well. In a paired in vitro study, we compared the quality of the final RBCs using HGM and LGM, and we determined whether differences in quality were related to differences in glycerol concentration or to the freeze-thaw process.

Materials and methods

Whole blood donations and processingWBC reduction of blood components to less than 1x106 WBCs per unit is mandatory in the Netherlands. WBC reduction by freeze-thaw-wash is around


28 29

95%,17,26,27 which is still above the limit.24,25 Therefore, we used filtered whole blood as starting material in this study.

The filtered units were subsequently pooled and split. After plasma removal, the units were either stored at 4⁰C in SAGM or treated with 40% or 19% glycerol (Fig. 1). The glycerolized RBCs were either immediately washed or frozen to -80°C or -196°C, respectively, and washed after thawing. We used a sterile connection device (model SC-201 AH, Terumo Europe, Leuven, Belgium) to perform all necessary tube welding. The washed cells were stored for 36 days at 4°C in SAGM.

In detail, 20 volunteers, who met AABB requirements for blood donors and provided informed consent, donated whole blood (500 mL, 66 ± 1 g of Hb) into a 600 mL bag (NPBI-Fresenius, Emmer-Compascuum, the Netherlands) containing 70 mL of CPD. Within 18 hours after collection and storage at room temperature, the units were WBC depleted (Imuflex, Whole Blood Filter, Terumo, Tokyo, Japan), which resulted in units containing 61 ± 2 g of Hb (a loss of 5.4 ± 1.6 g of Hb). After WBC depletion, we pooled 5 units in a 2 L bag

(PL 1813/1, Baxter, Deerfield, IL) and, after sampling, split the pool into 5 equal units (58 ± 4 g of Hb), using 800 mL bags (PL 146, Baxter, Deerfield, IL). The units were centrifuged for 4 minutes at 1615 x g, supernatant plasma was removed, and aliquots of plasma were stored at -80°C for plasma stability tests (see below).

The units of RBCs were treated in five different ways (Fig. 1). Total processing time took about 22 hours at room temperature from the time of donation until storage at the indicated temperatures. One unit (WBC depleted, Hct ∼ 0.80) was diluted with SAGM (Haemonetics) to an Hct of approximately 0.43, split into 8 parts of 30 mL in PVC bags (100 mL, Compoflex, NPBI-Fresenius, Emmer Compascuum, the Netherlands), and subsequently stored at 4°C. The four other units of the pool were treated with either 40% or 19% glycerol. The cells treated with glycerol were either directly deglycerolized as described below, using an automated closed washing system (ACP 215, Haemonetics) or frozen to -80°C or -196°C, respectively. After being frozen for 4 to 6 weeks, we thawed and deglycerolized these units. Approximately 4 g of Hb was lost due to glycerolization, sampling, bag transfer, and removal of supernatant glycerol. On average, 54 ± 4 g of Hb was recovered in the waste bag and component bag after thawing and deglycerolization with the closed washing system. The cells were resuspended in SAGM during the last deglycerolization step (Hct 0.50 ± 0.02, n = 16), split into 8 parts of 30 mL, and stored in PVC bags at 4°C. At different times during storage (as indicated in the figures), we took the PVC bags out of the refrigerator for sampling.

GlycerolizationBriefly, 38% wt/vol glycerol (containing 2.9% wt/vol sorbitol and 0.63% wt/vol NaCl, NPBI) was added manually in about 2 minutes to an equal volume of RBCs (Hct ∼ 0.80) to obtain a final concentration of 19% glycerol for LGM. For HGM, 57% wt/vol glycerol (containing 1.6% sodium lactate, 0.03% KCl, 0.0517% Na2HPO4, 0.1242% NaH2PO4, pH 6.8; Baxter) was added with the closed washing system in about 10 minutes to RBCs (Hct ∼ 0.80), proportionally to unit weight (a modifiable parameter of the ACP 215) to obtain a final concentration of 40% glycerol. Osmolality change during this process was kept at a constant rate of 500 mOsm per kg per minute.

Fig. 1. Schematic overview of study design

LeukoFilter

Filtered blood RBCsPlasma

19% glycerol (LGM)

DeglycerolizationACP215

(50 mL 6%NaCl1870 mL 0.9%NaCl)

RBC+SAGM4°C

control

Pool with 5 unitsfiltered blood;

split centrifuged

separation of plasma and RBCs

Whole Blooddonation

RBCs RBCs RBCs RBCs RBCs

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19% glycerol (LGM)


(50 mL 12%NaCl1870 ml 0.9%NaCl) Deglycerolization

ACP215(50 mL 6%NaCl

1870 mL 0.9%NaCl)


(50 mL 12%NaCl1870 mL 0.9%NaCl)

-196°C6 weeks

- 80°C6 weeks

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4°CLGM frozen

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Removal supernatantglycerol




30 31

Freezing and thawingThe LGM-treated cells (Hct ∼ 0.40) were transferred to an aluminum container (600 mL) (Boxal, Veenendaal, the Netherlands) and frozen within 10 minutes in liquid nitrogen.4,28 HGM-treated cells were centrifuged to remove excess glycerol (10 min at 1248 x g, final Hct ∼ 0.70). The PVC bag (Baxter) containing the RBCs was sealed in a plastic overwrap, placed in a cardboard box (Cekumed, Ooltgensplaat, the Netherlands), and put on the bottom of a -80°C freezer, inducing a slow drop in temperature (1-3°C/min) to -80°C.9 Thawing in a 37°C water bath took about 10 minutes and 20 minutes for LGM and HGM frozen RBCs, respectively. All units were thawed and warmed to a temperature of 30°C. Before deglycerolization, excess glycerol in the LGM units was removed by centrifugation at 2250 x g for 5 minutes.

DeglycerolizationThe closed washing system (ACP 215) was developed and FDA approved for the glycerolization and deglycerolization of HGM cells.24,25 For LGM, we developed a new deglycerolization procedure that would fit the programming of the closed washing system (ACP 215), washing fluids, and disposables (see below). For this new procedure, we compared the components obtained from the standard LGM procedure28 with the components obtained after introduction of the changes described below, using standard frozen LGM units.28 In the first washing step with hypertonic solution, 17.5% (wt/vol) sorbitol was replaced by a smaller volume of 6% (wt/vol) saline to make this step analogous to the addition of 12% (wt/vol) saline in HGM (see below) and to avoid volume overload in the bag. Secondly, we compared the current discontinuous washing procedure with a continuous washing procedure (carried out on an M 115 cell washer; Haemonetics Corp., Braintree, MA). These changes not only promoted recovery after the deglycerolization procedure (6% saline vs. 17.5% sorbitol) but also slightly shortened the processing time (continuous vs. discontinuous washing) and, hence, were considered to be appropriate changes to LGM (results not shown).

In the current study the standard programming of the closed washing system was used for deglycerolization of both LGM and HGM cells. For the deglycerolization of cells treated with 40% glycerol, 50 mL of 12% (wt/vol) saline was added to the glycerolized (and thawed) cells. For the deglycerolization of cells treated with 19% glycerol, 50 mL of 6% (wt/vol) saline was added. The

rest of the closed washing system deglycerolization procedure was completely the same for both LGM- and HGM-treated cells. After an incubation period of 2.5 minutes, 340 mL of 0.9% (wt/vol) saline and 0.2% (wt/vol) glucose was added. The cells were incubated for 1 minute. The diluted cells were subsequently pumped to the bowl, and supernatant saline was removed during centrifugation and pumped to the waste bag. The RBCs were returned to the original bag with an additional volume of a fresh 50 mL of 0.9% (wt/vol) saline and 0.2% (wt/vol) glucose to rinse the bowl. The cells were then diluted a second time with 400 mL of 0.9% (wt/vol) saline and 0.2% (wt/vol) glucose. After 1 minute of incubation, the diluted cells were pumped to the bowl and washed with 1080 mL of 0.9% (wt/vol) saline and 0.2% (wt/vol) glucose. In the final step, the cells were washed with, and resuspended in, 240 mL of SAGM and pumped to the storage bag. On average, the total closed washing system procedure of 1 unit took 65 ± 7 minutes (mean ± SD, n = 16 procedures).

Hb content and hemolysis during processingThe washing fluids collected in the waste bags and all storage and component bags were sampled to determine the total amount of Hb and the amount of Hb in supernatants of the samples after centrifugation for 10 minutes at 1400 x g. Hb was measured using the cyanmethemoglobin method.29 Hemolysis was expressed as a percentage of total Hb by using the following formula: Hemolysis = 100% x {(supernatant Hb) x (supernatant volume) / {(total Hb) x (total volume)}

Plasma stability and osmotic resistanceWe used two different concentrations of saline to estimate osmotic resistance. Samples (50 μL) at days 1 and 36 of units were diluted in 2 mL of 0.1% (wt/vol) Triton X-100 and 0.001% (wt/vol) saponin, 0.5% (wt/vol) saline, 0.6% (wt/vol) saline, 0.9% (wt/vol) saline or in autologous pool plasma. These samples were incubated for 30 minutes at 22°C and subsequently centrifuged at 1400 x g for 10 minutes. To meet the required increased sensitivity, Hb in the supernatant was measured directly on a spectrophotometer (Spectronic 301, Milton Roy, Ivyland, PA) using wavelengths 542 an 415 nm for samples in saline and 600, 577, 542, 510, 415, and 370 nm for samples in plasma.30 Hemolysis was calculated using the following formula:Hemolysis = 100% x (Hb sample) / (Hb supernatant in Triton/saponin)


32 33

RBC density distributionSamples of days 1 and 36 stored units were centrifuged at 1400 x g for 10 minutes, supernatant was removed, and 6 mL of RBCs were mixed with 25 mL of a Percoll solution (323 mOsmol/L, pH 8.24), slightly modified from Lutz et al.31 1 L of buffered Percoll solution contained 0.0372 g EDTA, 1.00 g D-glucose, 1.332 g NaCl, 0.356 g Na2HPO4, and 948 g Percoll (Pharmacia, Uppsala, Sweden).

The Percoll and RBCs were mixed and centrifuged at 43000 x g for 20 minutes at 16 to 23°C. The self-formed Percoll gradient with layers of RBCs was aspirated at a rate of 1 mL per minute from the bottom of the tube. Fractions of 3.5 mL were collected, and Hb content was measured in each fraction using the cyanmethemoglobin method.29

RBC metabolismSamples (0.5 mL) of the various RBC suspensions were diluted 1 in 1 in SAGM and deproteinized by adding 0.3 mL of 14% (wt/vol) perchloric acid.32,33 After 10 minutes on ice, samples were centrifuged and the protein-free supernatant was neutralized with ice-cold 2 N KOH and 0.2 mol MOPS per L. Neutralized extracts were subsequently stored at -80°C.

Adenine nucleotides were analyzed by high-performance liquid chromatography (HPLC), as described by De Korte et al.32,33 Lactate was determined enzymatically in these neutralized perchloric acid extracts by lactate oxidase and hydrogen peroxide activity (Sigma Chemical, St. Louis, MO). 2,3-DPG was measured enzymatically using the combined action of 2,3-DPG phosphatase, phosphoglycerate kinase, and glyceraldehyde-3-phosphate dehydrogenase according to the manufacturer’s instructions (Boehringer, Mannheim, Germany). Neutralized perchloric acid extracts were also used to determine the residual concentration of glycerol. Glycerol was measured enzymatically using the combined action of glycerokinase, pyruvate kinase, and lactate dehydrogenase (Diffchamb-Biocontrol, Nieuwerkerk aan den IJssel, the Netherlands).

Statistics

Data were analyzed using software (Microsoft Excel, Bellevue, WA) and the paired Student’s t-test. In general, results are depicted as means ± SD of the

number of observations given in parentheses. A p- value of less than 0.05 was considered significant.

Results

DeglycerolizationAs shown in Fig. 2, glycerolization of RBC units followed by deglycerolization without freezing already caused significant hemolysis. LGM cells showed considerably less hemolysis (2.4 ± 0.4%, n = 4) during deglycerolization than HGM cells (11 ± 3%, n = 4). The freeze-thaw process did not further affect hemolysis during deglycerolization of HGM RBCs (10 ± 2%, n = 4), whereas it almost doubled hemolysis during deglycerolization of LGM RBCs (4.4 ± 1.0%, n = 4). Clearly, total hemolysis during washing was significantly lower when LGM was used as compared to HGM.

Fig. 2. Hemolysis during deglycerolization. Hb content of the waste fluid supernatant was measured and expressed as a percentage of the total Hb content of the thawed unit before deglycerolization. Q, LGM method; O, HGM method.

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34 35

The final Hb content of the deglycerolized LGM and HGM units showed no difference because this parameter was mainly determined by the (limited) bowl volume, which we found to be 170 mL of RBCs. Volumes of RBCs exceeding this limit lead inevitably to cell spillage. Cell spillage was highest with the LGM cells (7 ± 5%) compared to HGM cells (4 ± 3%). Consequently, all our procedures yielded deglycerolized components with a cellular Hb content exceeding 36 g per unit (48 ± 2 g of Hb/ unit, n = 16) and free Hb below 0.2 g per unit (0.13 ± 0.05 g/unit, n = 16) in conformance with European guidelines.22 The remaining glycerol concentrations were 0.02 ± 0.01% wt/vol in our LGM samples and 0.04 ± 0.01% wt/vol in the HGM samples, all well below the critical residual glycerol concentration of 1 to 2%.6,14,19,20 There was no difference between frozen and non-frozen units, and glycerol concentrations remained the same during storage (results not shown).

Storage after washingIn the final washing step, the cells were resuspended in SAGM. Immediately after processing, hemolysis did not differ between treatments (0.28 ± 0.10%, n = 16; Fig. 3). However, hemolysis was significantly higher in all LGM and HGM cells compared to the untreated control RBCs (0.11 ± 0.05%, n = 4; Fig. 3).

During storage from days 1 to 36, glycerol treatment without freezing increased hemolysis. This effect of glycerol on RBCs was similar with both glycerol concentrations (Fig. 3).

In frozen and thawed cells, however, a pronounced difference was observed between the two methods. From day 1 through day 8 of storage, hemolysis in LGM (-196°C) cells was significantly higher than in HGM (-80°C) cells (Fig. 3). This difference between the two freeze-thaw procedures disappeared after day 15 of storage.

Osmotic resistance and plasma stabilityTo evaluate osmotic RBC resistance after storage in SAGM, hemolysis was measured on days 1 and 36 of storage. On day 1, we hardly saw an effect of treatment, whereas on day 36, a reduction of osmotic resistance due to deglycerolization and freezing and thawing was observed. Both on day 1 and day 36, hemolysis in plasma was 60 percent lower than in 0.9% saline.

No differences were observed between HGM and LGM cells.

RBC density distributionHemolysis during deglycerolization was relatively high with HGM RBCs (Fig. 2) but relatively low during storage after thawing (Fig. 3). The higher stability after thawing of frozen and thawed HGM cells may have been due to selection of a population of relatively strong (and young) cells. Differences in RBC density have been attributed to RBC aging.31 We therefore studied cell density on days 1 and 36 of storage using a continuous Percoll gradient.31 No significant change in density distribution was noted among any of the suspensions tested. Compared to day 1 of storage, we did measure a significant increase of cells in the highest

Fig. 3. Hemolysis during storage in SAGM after wash. Component was stored for 35 days at 4°C in SAGM. On the indicated time points, the Hb content of the component supernatant was measured and expressed as a percentage of the total Hb content of the component. F, control cells; , LGM non-frozen cells; , LGM frozen cells; G, HGM non-frozen cells; O , HGM frozen cells.


36 37

density fraction of cells after day 36 of storage (from 0.7 to 1.4%), but, again, there was no significant difference between treatments. This indicates that, if these high-density cells are aged RBCs, we could not detect selective removal of these older cells.

RBC metabolismTo explain the lower stability after thawing of LGM cells, several parameters of RBC metabolism were measured. The 2,3-DPG content of the control cells was already very low after day 1 of storage (3.9 ± 1.1 μmol/g of Hb, n = 4) and below the detection limit after day 2 of storage. The 2,3-DPG content appeared to be somewhat lower in the glycerol-treated cells, but these differences were not significant (results not shown).

Production of lactate did not differ between treatments during the first 8 days of storage, but from day 15 the control cells showed a significantly higher lactate production (on day 15 control cells 90 ± 12 μmol/g of Hb vs. glycerol treated cells 61 ± 9 μmol/g of Hb).

As shown in Fig. 4A the ATP content of the cells was not significantly affected by the treatments during the first two days of storage. Thereafter, ATP was significantly lower in the glycerol-treated cells. As was observed for lactate production, there was no difference among the various treatments. We also measured ADP and AMP to determine the total adenylate energy charge of the stored cells, because this parameter shows a better correlation with survival than just ATP.34 The ADP content and AMP content were significantly higher in the glycerol treated cells from day 1 on. There was no difference in the ADP content between the treatments, whereas the AMP content was significantly higher in the LGM and HGM frozen cells from day 8 on. From these data the total adenylate charge (ATP + 0.5 ADP)/(ATP + ADP + AMP) was calculated, which showed that the glycerol-treated cells had a lower net energy charge from day 1 on (Fig. 4B). On day 15, the frozen cells showed a significantly lower energy charge as compared to the non-frozen cells. None of these parameters showed a significant difference between LGM and HGM cells.

Fig. 4. Energy content during storage in SAGM after wash. Component was stored for 35 days at 4°C in SAGM. On the indicated time points, the ATP, ADP, and AMP content of the cells was measured, expressed as μmol per g of Hb, and from this the adenylate charge was calculated. (A) ATP content; (B) adenylate charge. F, control cells; , LGM non-frozen cells;, LGM frozen cells; G , HGM non-frozen cells; O , HGM frozen cells.

0 2 4 6 8 10 12 14 160 2 4 6 8 10 12 14 16

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38 39

Discussion

In this study, we compared two established methods for the cryopreservation of RBC using the newly developed washer (ACP 215) for deglycerolization. With the development of this fully automated and functionally closed washing system, it is now possible to deglycerolize RBCs while maintaining sterility, allowing subsequent storage beyond 24 hours.23-25 We studied the application of this washer for RBCs cryopreserved with LGM, and we compared the stability of these cells (during washing and subsequent storage) with RBCs prepared according to HGM.

Our study showed that by applying LGM, little hemolysis occurred during deglycerolization on the closed washing system and that freezing almost doubled this hemolysis. In contrast, the reverse was observed in HGM cells, in that more Hb was lost due to hemolysis during deglycerolization, whereas freezing hardly had an additional effect (Fig. 2). Hemolysis immediately after thaw and wash was similar for both LGM and HGM cells with or without freezing, showing that all free Hb was removed by the closed washer system deglycerolization procedure. Though relatively small numbers of units were used for the current study, the component loss due to hemolysis we observed during the LGM and HGM procedures is comparable to the losses (HGM 10-20%, LGM 5-10%) reported by others using the same24,25 or different deglycerolization procedures.5,9,15,17,24-28,35-38 All processed units met US and European guidelines for thawed deglycerolized RBCs.21,22 From these results it can be concluded that the closed washing system (ACP 215) can successfully be applied, not only to HGM, but also to LGM. However, to fully exploit the advantage of the washer to LGM, a liquid nitrogen resistant cryopreservation bag, suitable for sterile docking onto the washing device, is required to enable prolonged storage after washing beyond 24 hours.

Interestingly, the decreased stability of RBCs during washing caused by the higher glycerol concentration was not reflected in a decreased stability during storage after washing. Actually, during the first week of storage, hemolysis was highest for LGM frozen cells (Fig. 3). This lower stability after thawing of LGM cells was not due to the processing on the closed washing system because we observed similar differences in stability when processed by our current routine procedures.39 Two alternative explanations were considered to account for the lower stability after thawing of LGM cells. Firstly, the higher hemolysis during storage of LGM cells could be due to the preservation of a more labile (and

aged) population of RBCs. However, the density distributions of the different concentrates did not support this hypothesis. Alternatively, the lower hemolysis of HGM cells during storage after thawing could reflect a better protection against cell damage induced by the freeze and thaw process. This is indirectly supported by our finding that hemolysis in HGM cells during washing and during the first week of storage was not affected by freezing. Furthermore, hemolysis of HGM frozen cells was lower during storage as compared to LGM frozen cells. On the other hand, this improved protection by high glycerol leads to a higher degree of hemolysis during washing, most probably inflicted by a higher degree of osmotic changes as compared to LGM.

Other than hemolysis, no other parameter studied showed differences between the frozen HGM and the frozen LGM cells within the first week of storage. In contrast to previously published reports,14,40 we observed strongly reduced 2,3-DPG levels in all concentrates already at day 1. It was previously recognized that holding fresh whole blood for 24 hours at room temperature reduces 2,3-DPG levels from 13.1 to 4.4 μmol per g of Hb.41 In our experiments, the average holding time at room temperature between donation and time of storage was 22 hours. This may explain the above-mentioned 2,3-DPG levels.

In accord with data from Moroff and Meryman,42 we demonstrated that all glycerolization and deglycerolization procedures with or without freezing resulted in a final product with a normal ATP content per cell and that this ATP level was maintained during the first 2 days of storage (Fig. 4A). However, we also showed that the adenylate content of the cells on the first day of storage was significantly lowered by the various treatments (Fig. 4B). Because the total adenylate content shows a better correlation with survival than just ATP,34 this may predict a lower “in vivo” survival of these SAGM-stored thawed and washed cells.

Similar to other investigators, we have found that, compared to SAGM, both previously frozen HGM and LGM RBCs demonstrated a reduced hemolysis in AS-3.23,37,39 “In vivo” recovery of the closed washing system-processed HGM cells, stored for 15 days in AS-3, appeared to be satisfactory;24,25 for LGM cells, this remains to be shown.


40 41

Acknowledgments

The authors thank Dirk de Korte, PhD, and Eric Gouwerok (Sanquin Research at CLB, Amsterdam, the Netherlands) for their help with adenine nucleotide determinations.

References


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15. Moss GS. Preservation of red cells by freezing. Surg.Annu. 1970;2(0):35-50.16. Chaplin HJ. Frozen red cells revisited. N.Engl.J.Med 1984;311(26):1696-8.17. Crowley JP, Wade PH, Wish C, Valeri CR. The purification of red cells for transfusion

by freeze-preservation and washing. V. Red cell recovery and residual leukocytes after freeze-preservation with high concentrations of glycerol and washing in various systems. Transfusion 1977;17(1):1-7.

18. Chaplin HJ. The proper use of previously frozen red blood cells for transfusion. Blood 1982;59(6):1118-20.


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19. Huggins C. Preparation and usefulness of frozen blood. Annu.Rev.Med 1985;36:499-503.

20. Valeri CR. Frozen blood (concluded). N.Engl.J.Med. 1966;275(8):425-31.21. Standards for Blood Banks and Transfusion Services, 20th edition, Bethesda:

AABB, 2000. 22. Guide to the Preparation, Use and Quality Assurance of Blood Components, 7th

edition, Strasbourg: Council of Europe, 200123. Hess JR, Hill HR, Oliver CK, Lippert LE, Greenwalt TJ. The effect of two additive

solutions on the postthaw storage of RBCs. Transfusion 2001;41(7):923-7.24. Valeri CR, Ragno G, Pivacek LE, Srey R, Hess JR, Lippert LE, Mettille F, Fahie

R, O’Neill EM, Szymanski IO. A multicenter study of in vitro and in vivo values in human RBCs frozen with 40-percent (wt/vol) glycerol and stored after deglycerolization for 15 days at 4 degrees C in AS-3: assessment of RBC processing in the ACP 215. Transfusion 2001;41(7):933-9.

25. Valeri CR, Ragno G, Pivacek L, O’Neill EM. In vivo survival of apheresis RBCs, frozen with 40-percent (wt/vol) glycerol, deglycerolized in the ACP 215, and stored at 4 degrees C in AS-3 for up to 21 days. Transfusion 2001;41(7):928-32.

26. Crowley JP, Skrabut EM, Valeri CR. Immunocompetent lymphocytes in previously frozen washed red cells. Vox Sang. 1974;26(6):513-7.

27. Crowley JP, Valeri CR. The purification of red cells of transfusion by freeze preservation and washing. II. The residual leukocytes, platelets, and plasma in washed,freeze-preserved red cells. Transfusion 1974;14(3):196-202.

28. Krijnen HW, Kuivenhoven AC, De Wit JJ. The preservation of blood cells in the frozen state. Experiences and current methods in the Netherlands. Cryobiology 1968;5(2):136-43.

29. Zijlstra WG, Van Kampen E. Standardization of hemoglobinometry. I. The extinction coefficient of hemiglobincyanide. Clin.Chim.Acta 1960;5:719-26.

30. Vogelaar EF, Brummelhuis HG, Beentjes SP, Krijnen HW. Contributions to the optimal use of human blood. I. Analysis and optimalization of the production of plasma protein fraction (PPF). Vox Sang. 1972;23(6):481-92.

31. Lutz HU, Stammler P, Fasler S, Ingold M, Fehr J. Density separation of human red blood cells on self forming Percoll gradients: correlation with cell age. Biochim.Biophys.Acta 1992;1116(1):1-10.

32. de Korte D, Haverkort WA, Roos D, van Gennip AH. Anion-exchange high performance liquid chromatography method for the quantitation of nucleotides in human blood cells. Clin.Chim.Acta 1985;148(3):185-96.

33. de Korte D, Haverkort WA, van Gennip AH, Roos D. Nucleotide profiles of normal human blood cells determined by high-performance liquid chromatography. Anal.Biochem. 1985;147(1):197-209.

34. Hogman CF, de Verdier CH, Ericson A, Hedlund K, Sandhagen B. Studies on the mechanism of human red cell loss of viability during storage at +4 degrees C in vitro. I. Cell shape and total adenylate concentration as determinant factors for posttransfusion survival. Vox Sang. 1985;48(5):257-68.

35. Valeri CR, Pivacek LE, Cassidy GP, Ragno G. Posttransfusion survival (24-hour) and hemolysis of previously frozen, deglycerolized RBCs after storage at 4 degrees C for up to 14 days in sodium chloride alone or sodium chloride supplemented with additive solutions. Transfusion 2000;40(11):1337-40.

36. Valeri CR, Pivacek LE, Cassidy GP, Ragno G. The survival, function, and hemolysis of human RBCs stored at 4 degrees C in additive solution (AS-1, AS-3, or AS-5) for 42 days and then biochemically modified, frozen, thawed, washed, and stored at 4 degrees C in sodium chloride and glucose solution for 24 hours. Transfusion 2000;40(11):1341-5.

37. Moore GL, Ledford ME, Mathewson PJ, Hankins DJ, Shah SB. Post-thaw storage at 4 degrees C of previously frozen red cells with retention of 2,3-DPG. Vox Sang. 1987;53(1):15-8.

38. Derrick JB, McConn R, Sorovacu ML, Rowe AW. Studies of the metabolic integrity of human red blood cells after cryopreservation. II. Effects of low-glycerol-rapid-freeze preservation on glycolysis. Transfusion 1972;12(6):400-4.

39. Noorman F, Lelkens CCM, et al. Frozen red blood cells, comparison of the low glycerol and high glycerol method. Transfusion 2000;40:(Suppl):63S.

40. Rittmeyer IC, Nydegger UE. Influence of the cryoprotective agents glycerol and hydroxyethyl starch on red blood cell ATP and 2,3-diphosphoglyceric acid levels. Vox Sang. 1992;62(3):141-5.

41. Pietersz RN, de Korte D, Reesink HW, Dekker WJ, van den Ende A, LOOS JA. Storage of whole blood for up to 24 hours at ambient temperature prior to component preparation. Vox Sang. 1989;56(3):145-50.

42. Moroff G, Meryman HT. Influence of glygerol on ATP and 2,3-DPG levels of human erythrocytes. Vox Sang. 1979;36(4):244-51.

Charles C.M. Lelkens, Jack G. Koning, Bob de Kort, Ingeborg B.G. Floot and Femke Noorman

Military Blood Bank, Leiden, the Netherlands

Transfus Apher Sci. 2006 Jun; 34(3):289-98

Chapter

Experiences with frozen blood products in the Netherlands military

3

Chapter 3 Experiences with frozen blood products in the Netherlands military

46 47

Abstract

For peacekeeping and peace enforcing missions abroad the Netherlands Armed Forces decided to use universal donor frozen blood products in addition to liquid products. This article describes our experiences with the frozen blood inventory, with special attention to quality control. It is shown that all thawed (washed) blood products are in compliance with international regulations and guidelines. By means of the -80°C frozen stock of red cells, plasma and platelets readily available after thaw (and wash), we can now safely reduce shipments and abandon the backup “walking blood bank”, without compromising the availability of blood products in theater.

Introduction

Blood supply in the Netherlands is a responsibility of the Sanquin Blood Foundation. From the four regional, main blood banks - Northwest, Northeast, Southwest and Southeast - all Dutch hospitals are provided with the necessary blood products, derived from volunteer, unpaid donors. Sanquin operates nationwide in an environment in which demand and supply are more or less balanced. Furthermore, logistical problems are almost non-existent, given the size of the country and its infrastructure. Therefore, the usual shelf lives of particularly red cells and platelets hardly create a problem for the civilian community. The military system by contrast, operates in a totally different environment.

The Military Blood Bank (MBB), the first link in the Netherlands military blood supply chain, is dependent upon the civilian population in that its donors provide indirectly - through Sanquin - the blood products needed during deployments. All blood products are tested and processed by Sanquin, including those used for military purposes abroad. The MBB is responsible for providing the necessary blood products to deployed military expeditionary units abroad. Finding a balance between demand and supply as observed in the civilian community is virtually impossible, given the current nature and risks of military deployments.

In general, transfusing blood in the military situation will be related to trauma, because of massive blood loss and hypovolemic shock.1 In addition to volume replacement, casualties who have lost more than 25-30% of their original blood volume also require restoration of their oxygen-carrying capacity, which calls for transfusing red cells. At the same time, concurring coagulation defects will have to be countered by administering plasma and/or platelets.2,3 To exclude inadvertent clerical errors as much as possible and to reduce the number of units to be stockpiled, the use of universal donor products is imperative, i.e. O Rh (D) positive and negative red cells, O Rh (D) positive and negative platelets and AB plasma.

Blood products have a finite life span; the shelf life of the usual, standard liquid stored red cells is measured in weeks and for platelets even in days. If those products were to be readily available in theater, it would require shipments of platelets every week and red cells preferably every two weeks, but at least every


48 49

month. This limits their use and creates a substantial logistical burden, particularly, if increased numbers of blood units are needed at short notice. Newly developed additive solutions that significantly prolong storage times of red cells,4 would diminish the frequency of shipments to a certain extent, but these solutions are not available on the market yet. Newly developed storage solutions for platelets do not extend storage times beyond 10 days.5 Products like hemoglobin based oxygen carriers6 and artificial platelets5 are not yet available as an alternative.

In 1991, during the first Gulf War, the Netherlands Military Blood Bank shipped universal O (Rh D positive and negative) liquid RBC from Amsterdam to the United Arab Emirates, in support of a land-based Royal Netherlands Naval surgical team. Later on, in 1992, we shipped universal liquid RBC and fresh frozen plasma (FFP) to our medical treatment facilities (MTF’s) deployed in Cambodia and Bosnia. Platelets were provided by a so-called “walking blood bank”. The number of new deployments grew steadily and so did the pressure on the blood supply system. Providing standard blood products proved to be cost-ineffective and, moreover, could not guarantee the availability of blood products at all times. To date, freezing is the only technique available to substantially extend the shelf lives of the products we need and to reduce shipments to an absolute minimum. Deep frozen (-80°C) red cells can be stored for 10 years at least,7,8 deep frozen plasma for 7 years9 and deep frozen platelets for 2 years.10,11

This report describes the experiences of the Netherlands military with these frozen blood products, with special attention to quality control and compliance to (inter)national regulations. We show that it is now possible to deploy a self-sufficient military blood bank facility, based on an inventory of deep frozen red cells, plasma and platelets.


Red cellsWe procured leukodepleted, filtered whole blood (O, Rh D positive and negative) units from Sanquin Southwest (Rotterdam, the Netherlands), as raw material to produce the frozen red cells from. The unit was transferred to a 1 L PVC bag (MacoPharma, Tourcing, France) and centrifuged (Hettich Roto Silenta) at 1615 x g, 4 min, brake 0. The supernatant plasma was removed in a 600 mL PVC bag (Terumo, Tokyo, Japan) and a volume of 57% w/v glycerol (‘Glycerolite’

Baxter, Toronto, Canada) was sterilely added via the ACP215 (Haemonetics, Braintree, MA, USA) in about 11 min to a final concentration of around 40% (w/v). Subsequently the unit was centrifuged (Hettich Roto Silenta) at 1248 x g, 10 min, no brake, and the supernatant glycerol was removed in a 600 mL PVC Terumo bag. The final product bag was then folded and vacuum-sealed in a plastic overwrap bag. The bag was packed into a rigid cardboard box (Cekumed, Ooltgensplaat, the Netherlands) and frozen to a temperature of -80°C on the bottom of a mechanical freezer (Revco, PolyTemp Scientific, Bolsward, the Netherlands) resulting in a freezing rate of 1-3°C/min. After a minimum of 24 h, the product was placed in a quarantine inventory at -80°C (Revco). All units were frozen within 24 h after donation.

Prior to transfusion, the deep frozen erythrocyte concentrates (DEC) were thawed in a temperature controlled water bath (Forma Scientific, De Meern, the Netherlands), maintained at 42°C. When the unit reached a temperature of 30-35°C the unit was deglycerolized in the ACP 215, a semi-automated, functionally closed washing system. In a number of washing steps, the cells were sterilely washed with NaCl 12% (Baxter, Deerfield, IL, USA), a mixture of normal saline and 0.2% glucose (Baxter, Deerfield, IL, USA) and finally suspended in the storage solution AS-3 (Haemonetics, Braintree, MA, USA). The thawed, washed red cells were stored for 14 days at 2-6°C. Total processing time was 100-120 min after removal from the freezer.

PlasmaWe procured leukodepleted FFP (AB-Rh D positive and negative), isolated by apheresis, from Sanquin Southwest after ±1 year of quarantine storage at -30°C, and release from quarantine. Units were thawed in a temperature controlled water bath (Forma Scientific), maintained at 37°C, and warmed to a temperature of 20-30°C. The thawed plasma was then transferred to a 600 mL PVC bag (MacoPharma), folded and vacuum-sealed in a plastic overwrap bag. The unit was packed into a rigid cardboard box (Cekumed) and frozen to a temperature of -80°C on the bottom of a mechanical freezer (Revco) resulting in a freezing rate of 1-3°C/min. Before transfusion, the deep frozen plasma (DFP) units were thawed in 25-35 min in a temperature controlled water bath (Forma Scientific), maintained at 37°C, to 30-35°C. Total processing time was 25-35 min after removal from the freezer.


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PlateletsWe procured fresh, leukodepleted platelet concentrates, isolated by apheresis (Amicus), from Sanquin Southwest. Units were stored at 22°C with agitation for 12-20 h, until the freezing procedure was started. A transfer set (Baxter) was sterilely connected to the unit to be frozen and the unit was then put on a flatbed shaker (GFL, Burgwedel, Germany). A bottle, containing 27% (w/v) DMSO in 0.9% saline (Pharmacy Department, Central Military Hospital, Utrecht, the Netherlands) was hung at 7 in. in a Laminar Air Flow cabinet (PMV, Woerden, the Netherlands). The Teflon cap of the bottle was spiked with the transfer set. While the unit was rotating at 180 rotations per minute on a flatbed shaker (Beun De Ronde, Abcoude, the Netherlands), 75 mL DMSO was added by means of gravity (± 10 ml/min) in ± 7 min. The unit was subsequently transferred to a 400 mL PVC bag (Baxter) and centrifuged at 1250 x g for 10 min, brake 0. Supernatant DMSO-plasma was removed and the remaining 10-20 mL platelet concentrate was carefully mixed in the 400 mL bag by gently rubbing the bag with a nylon gauze. The platelet bag was folded, vacuum-sealed in a plastic overwrap bag, packed into a rigid cardboard box (Cekumed) and frozen to a temperature of -80°C on the bottom of a mechanical freezer (Harris) resulting in a freezing rate of 3-5°C/min. All units were frozen within 24 h of donation. Prior to transfusion, the DTC were warmed to 30-35°C in ± 5 min in a temperature controlled water bath (Forma Scientific) maintained at 37°C. After gently mixing the platelet unit by means of gently rubbing the bag with a nylon gauze, a thawed DFP unit was sterilely connected (TSCD, Terumo Europe, Leuven, Belgium) to the platelet bag. The thawed plasma was added to the thawed platelets and the platelets were easily suspended in the plasma by transferring the platelets in plasma back and forth to the platelet freezing bag. Hereafter, the platelets are ready for transfusion. Total processing time after removal from the freezer is 30-40 min.

Labels and logisticsAll blood products used were labeled conforming to ISBT guidelines. For frozen red cells and platelets an extra label for the thawed product was enclosed in the vacuum-sealed plastic overwrap bag. The frozen products were stored at -80°C in mechanical freezers, equipped with online alarm systems and CO2 back up, to minimize the consequences of unforeseen power outages, possibly leading to out-of-range inside temperatures. Units were transported in “dry ice” (ca. -80°C) in insulated small shipping containers with a maximum of 10 units each

(Dometic, Alphen aan den Rijn, the Netherlands) or bigger versions with a maximum capacity of 192 units (Olivo, Roche-la-Molière, France). Temperature during transportation was monitored continuously by a TempTale® device (TDS, Sassenheim, the Netherlands) in each container and never exceeded the upper limit of -65°C. Products were transferred to a -80°C freezer (Revco) in a room temperature controlled container. The freezer was connected to an audible and visible alarm system and CO2 back up. Temperature was monitored continuously with a 6 in. chart recorder on the freezer. All in-theater storage, thawing and washing procedures were performed in a room temperature controlled blood bank container, designed by the Royal Netherlands Army.

Quality control and analysesSanquin Southwest performed the quality control of donors, donations and leukodepletion. To check the final content of frozen/thawed products, a clinical hematology analyzer (Sysmex KX-21, Goffin Meyvis, Etten-Leur, the Netherlands) was used. For additional quality control of the RBC, capillary hematocrit and supernatant free hemoglobin (after centrifugation at 2200 x g, 10 min) was measured on a Plasma low hemoglobin device (HemoCue, Oisterwijk, the Netherlands). The percentage of hemolysis was determined by the ratio of free Hb to total Hb. We used a Reflotron® triglyceride test to measure the residual glycerol concentration in the supernatant after a 1:10 dilution in 0.9% saline. The KC4A (Amelung Coagulometer, Germany) was used to determine the concentration of coagulation factors V and VIII, APTT and PT, according to the manufacturer’s instructions. For these tests standard -80°C plasma, factor VIII deficient plasma and factor V deficient plasma from Cryocheck (Precision Biologic, Darmouth, NS, Canada) were used. pH was measured by means of an electrode (Delta OHM, Caselle di Sevazzano, Italy). All data were stored and analyzed in Excel 97. The differences were studied with the Student’s t-test; a p value < 0.05 was considered statistically significant.

Results

Leukodepleted RBC units should contain less than 1 x 106 WBC and less than 15 x 109 platelets. Liquid stored red cells should contain at least 40 g of Hb and hemolysis at the end of the storage period should not exceed 0.8% or 1%.6,10,12


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After deglycerolization, the final glycerol concentration should be less than 1% in order to prevent hemolysis in the recipient.13 For deglycerolized, stored red cells hemolysis should not exceed 1% at the end of the storage period.12

From March 2003 to July 2005, a total of 1360 units of filtered, leukodepleted units of whole blood have been processed to frozen red cells. Table 1 shows an overview of the characteristics of these units. From unit hematocrit prior to freezing (fixed on 80%) and the unit weight (determined for each unit) the ACP 215 accurately calculates the amount of glycerol needed for each individual unit to reach a final concentration of 40% w/v (40 ± 0.5%). Throughout the freezing procedure, only 3 ± 1 g Hb was lost due to sampling and removal of supernatant plasma and glycerol.

Units were deglycerolized, stored at 2-6°C for 14 days, after which the product quality was assessed. The process was fully validated in the Netherlands prior to in-theater validation and implementation. In April 2004, the thaw-wash process was validated in Iraq in the operational, military environment. For two days during 12 hours each day, two skilled lab technicians, using two ACP 215 devices, thawed and washed 22 units of DEC. There was no difference between the units processed at the MBB in Leiden and those in Iraq. At the end of April 2004, we therefore decided to implement the ACP 215 to deglycerolize units of DEC in Iraq. The backup system proved to be very useful when during the Iraq election period, logistics were hampered and no liquid red cells could be transported. Twenty units were thawed, washed and stored at 2-6°C successfully to replace the outdated liquid inventory without further delay.

An overview of the quality of thawed, deglycerolized and subsequently stored units (including the data from the 42 units processed in Iraq) is shown in Table 1. The ACP215 effectively removed glycerol from the thawed product from 40% w/v to below the 1% w/v upper limit.13 It appeared that the Hb content of the frozen unit was primarily dependent upon the Hb content of the leukodepleted product (Fig. 1A). The Hb content of the washed unit was primarily dependent upon the bowl size (275 mL), since hemolysis during washing (19 ± 5%) was independent of the Hb content of the frozen product, whereas cell spillage was not (results not shown). As shown in Fig. 1B, the Hb content of a unit before freezing should at least be 50 g in order to obtain the minimum required product content of 40 g of Hb after deglycerolization. Only 16 of 1360 units (1.2%) did not meet this minimum requirement of 50 g of Hb prior to freezing and these units were quarantined. Of these low Hb units, 14 units were washed and Hb

content was below 40 g of Hb in all cases (Fig. 1B). Various other reasons, such as training and education purposes (1.1%) and bad seals of the sterile connection device (1%), accounted for another 3.6% that were not released for transfusion. During the validation period 168 units were deglycerolized and 7 units (4.2%) were discarded. Product loss was caused by power failure (2 units), bowl damage during deglycerolization (2 units), ACP215 technical problems (1 unit) and leakage of the sterile connection (1 unit). Another unit was lost due to wrongfully connecting the washing fluids on the ACP215.

In conclusion, out of 1360 units, 1298 units (95.4%) met all the criteria to be stored at -80°C for a maximum of 10 years, ready to be used on deployments.

Table 1 Characteristics of deep frozen red cells before, during and after freezing to -80°C

Mean ± SD Limit(s) % OKFiltered whole blood (n = 1360)Thrombocytes (109/U) 2 ± 2 <15c 99.8White blood cells (106/U) 0.3 ± 0.2 <1a 99.6Volume (mL) 519 ± 13 470-570b 99.6Hb content (g) 63 ± 5 >55b 93.3

Deep frozen red cells (n = 1360)Hematocrit (L/L) 0.60 ± 0.03 0.50-0.70b 98.5Glycerol concentration (g/dL) 40.0 ± 0.5 38-42b 99.6Volume (mL) 326 ± 33 250-450b 99.6Hb content (g) 60 ± 5 >50b 98.8

Thawed washed red cells (n = 147)Hematocrit (L/L) 0.56 ± 0.03 0.50-0.65c 95.2Glycerol concentration (g/dL) 0.18 ± 0.03 <1.0d 100Volume (mL) 294 ± 4 >245c 100Hb content (g) 45 ± 3 >40a 95.2Hemolysis at day 14 (%) 0.6 ± 0.2 <1.0d 96.3Hemolysis at day 14 (%) 0.6 ± 0.2 <0.8a 87.7

a CE guidelines leukodepleted red cells.10

b MBB internal guidelines.c Sanquin internal guidelines leukodepleted red cells.d FDA/AABB guidelines deglycerolized red cells.4,12,13


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We found that filtered, leukodepleted whole blood, can effectively be processed with the ACP215 to produce DEC. The thawed washed products contained at least 40 g of Hb and can be stored for 14 days at 4°C with low hemolysis after deglycerolization and suspension in AS-3.

Table 2 shows that AB plasma has a high factor VIII and factor V content, even after storage for almost 1 year at -30°C. Compared to FFP a slight reduction of factor VIII concentration (p > 0.05) and factor V (p < 0.05) was observed

Table 2 Characteristics of deep frozen plasma before, during and after freezing to -80°C

Mean ± SD Limit(s) % OKFresh frozen plasma apheresis, leukodepleted (FFP)Thrombocytes concentration (109/L) (n = 68) 5 ± 7 <50a 100White blood cells (106/U) No datad <1c >90Volume (mL) (n = 188) 290 ± 15 225-315c 99.5

Coagulation after FFP storage, 0.9 ± 0.01 year at -30°CAPTT (s) (n = 12) 30.0 ± 2.2 <36b 100PT (s) (n = 12) 12.2 ± 0.5 <14b 100Factor VIII (U/mL) (n = 12) 1.4 ± 0.5 >0.70a 100Factor V (U/mL) (n = 12) 1.0 ± 0.2 >0.70b 100

Deep frozen plasma (DFP)Thrombocytes concentration (109/L) (n = 27) 10 ± 8 <50b 100Volume (mL) (n = 188) 284 ± 14 225-315b 99.5

Coagulation after DFP storage, 0.76 ± 0.02 year at -30°C, followed by 1.31 ± 0.02 years at -80°C

APTT (s) (n = 12) 31.0 ± 2.3 <36b 100PT (s) (n = 12) 13.0 ± 0.5 <14b 100Factor VIII (U/mL) (n = 12) 1.2 ± 0.3 >0.70b 100Factor V (U/mL) (n = 12) 0.9 ± 0.1 >0.70b 100

a CE guidelines fresh frozen plasma (apheresis).10

b MBB internal guidelines.c Sanquin internal guidelines fresh frozen plasma apheresis, leukodepleted.d Product is manufactured by Sanquin Southwest, according to Sanquin internal guide-lines.

Fig. 1. The relation between Hb content of ground substance and Hb content of final product. (A) The relation between Hb content of leukodepleted whole blood and Hb content of frozen red cells (N = 1360). (B) The relation between Hb content of frozen red cells and Hb content of thawed, washed red cells (N = 161).

85

80

75

70

65

60

55

50

45

40

A

5250484644424038363432302826

B

40 50 60 70 80 90Hb content of leukodepleted whole blood (gram Hb)

Hb content of frozen product (gram Hb)40 45 50 55 60 65 70 75 80

Hb

cont

ent o

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uct (

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, was

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in DFP. The concentration of factors V and VIII did not decrease to below the minimum of 70%. The volume of plasma is reduced by ± 6 mL because of sampling, without reducing the plasma volume below the minimum of 225 mL. Thus, thawing and subsequent freezing of AB plasma to -80°C can be performed, without deviation of the product quality from the Council of Europe (CE) guidelines.10 Until now, a total of 435 DFP units were produced and only 16

Table 3 Characteristics of deep frozen platelets before, during and after freezing to -80°C

Mean ± SD Limit(s) % OKPlatelets apheresis, leukodepleted (n = 217)Thrombocytes concentration (109/L) 1221 ± 220 <1500a 90.2White blood cells (106/U) 0.1 ± 0.1 <1a 100Volume (mL) 312 ± 19 150-400c 99.5Thrombocytes (109/U) 381 ± 68 >200a 100pH 7.1 ± 0.1 >6.8c 100

Frozen platelets (n = 217)Volume (ml/U) 14 ± 4 <20b 92.6Thrombocytes (109/U) 349 ± 58 >220b 98.2Platelet recovery start product–frozen product (%) 93 ± 13 >80b 97.2DMSO concentration (g/dL) 5.1 ± 0.4 4-6b 97.7DMSO content (g) 0.7 ± 0.2 <1b 94.0

Thawed resuspended platelets in DFP (n = 27)Platelet recovery start product-frozen product (%) 94 ± 15 >80b 92.6Platelet recovery freeze-thaw 83 ± 17 >40d 100Platelet recovery start product-end product 77 ± 15 >40b 100Thrombocytes concentration (109/L) 977 ± 226 <1500a 92.6Volume (mL) 279 ± 28 225-315b 96.3Thrombocytes (109/U) 269 ± 45 >200a 100DMSO content (g) 0.7± 0.2 <10b (d) 100pH 7.7± 0.1 >6.8c 100

a CE guidelines fresh frozen platelets apheresis, leukodepleted.10

b MBB internal guidelines.c Sanquin internal guidelines fresh frozen platelets apheresis, leukodepleted.d CE guidelines cryopreserved platelets.10

units (3.7%) were discarded. The majority (12 units) was discarded because the units were thawed in a defective water bath, which resulted in too slow warming of the units. Furthermore, two units were discarded due to a high RBC content and another two units due to the use of spiking because of short tubing on the original FFP bag.

An additional advantage of the DFP procedure is that the folding of the product bag and packaging in a vacuum-sealed overwrap bag, in combination with the rigid cardboard box, prevents breakage of the PVC bag. We found 8.2% breakage when thawing -30°C stored FFP (n = 486), and no breakage at all with DFP (n = 82). Similarly, the above-mentioned thawed DEC (n = 167) also did not show any breakage, nor did the DTC (n = 27) described below.

Thawed DTC units were suspended in thawed DFP and the amount of platelets was determined. As shown in Table 3, the units contained more than the minimum of 200 x 109 platelets/unit (CE guidelines10 for fresh liquid platelets) and showed a higher than 40% recovery (CE guidelines10 for frozen platelets). The pH was well above 6.8, varying between 7.6 and 7.8. In the CE guidelines10

a limit is set to the volume of product that can be transfused without washing. We concluded from these guidelines that no more than 10 g of DMSO (200 mL 5% w/v) should be present in one platelet unit. As shown in Table 3, the DMSO content of the unit is very low and did not exceed this limit; in fact 94% of all units have a DMSO content below 1 g/unit. Out of the 217 units frozen, 13 units (6%) were discarded. This was due to recall because of a positive BacT/Alert® measurement (2 units), clerical or technical errors (4 units), presence of aggregates prior to freezing (2 units), and failure of a sterile connection device (4 units). Only 1 unit was discarded due to a large starting volume, resulting in a final DMSO concentration below 4%.

Discussion

It has been shown that the high-glycerol method can be used to freeze red cells with an acceptable postthaw and postwash recovery. By means of the ACP215 it is now possible to store the red cells after washing for a period of 14 days at 4°C with an acceptable postthaw hemolysis and “in vivo” survival.8,14-16 Platelets can be frozen in 5% DMSO at -80°C.17,18 Although in vivo survival is reduced,17-21 the frozen platelets are more effective in reducing blood loss, compared to three day


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stored liquid platelets.21 Fresh frozen plasma (-30°C) has been used worldwide; the effect of refreezing them to -80°C has not been studied before. To ensure readily available blood products on missions all over the world, the Netherlands Ministry of Defense decided to build a state-of-the art frozen inventory of the blood products most needed: red cells, plasma and platelets.

Red cells can be frozen in either a high (40% w/v) glycerol or a low (20% w/v) glycerol concentration. The 40% glycerol allows for storage at -80°C in a mechanical freezer, whereas the 20% glycerol requires liquid nitrogen (-196°C). We have shown that the -80°C frozen cells are more stable during postwash storage.22 Furthermore, introducing -80°C to freeze red cells fitted well into the concept of a deployable, integrated liquid-frozen blood bank.23 The feasibility of the concept itself had been shown before, as well as the use of previously frozen red cells in the treatment of combat casualties.24-26 Using the -80°C frozen RBC allows for 10 years of storage, although much longer periods have been shown to be feasible.7,8

To deglycerolize the thawed red cells in theater, prior to transfusion, we previously have used the M 115 (Haemonetics, Braintree, MA, USA), a non-closed washing device, permitting a 24 h postwash storage period only. Our frozen red cells thus could not serve as a primary source to meet our red cell needs “in the field”, but were used as a backup for the liquid stock, which was supplied every 2 weeks. We used this concept during missions in Bosnia (1993-2005), Afghanistan (2002-2003) and Liberia (2003). Both in Afghanistan and Liberia the frozen red cells have proven to be essential when sudden demand exceeded the available liquid stock of red cells.

With the introduction of the ACP215, a semi-automated, closed washing device, postthaw, postwash storage of frozen red cells became possible for at least 14 days at 2-6°C.8,14-16 We show that the ACP215 in its current, FDA approved, configuration can be used effectively to produce a stock of DEC and to produce on demand liquid red cells in theater with sufficient product yield and low hemolysis after storage. In addition, we show that leukodepleted whole blood collected in CPD can be used as a source for DEC.

Furthermore, in 2005 it became clear in Iraq that a military hospital blood bank facility can be deployed without regular shipments of liquid red cells. It is able to meet the needs of a surgical team by thawing and washing a certain number of frozen units once a week. This creates a new concept of an integrated liquid-frozen blood bank. We have successfully used this new concept in the

current mission in Afghanistan. Our conclusion is that the ACP215 enables a more efficient and flexible use of RBC for military deployments, still meeting the standard quality requirements of the regulatory civilian authorities.

It is well known that following massive blood replacement, plasma and platelet transfusion is required to correct coagulation defects. It has been shown that among survivors of massive blood transfusion more platelets were transfused.27 Fresh whole blood or stored platelet concentrates are required to correct the coagulation defect.3,27-29 During several missions in the early nineties, a so-called “walking blood bank”, consisting of voluntary military personnel in theater, was to provide whole blood, primarily as a source of platelets. In 1996, however, we encountered a case of hepatitis B transmission in Bosnia, derived from a “walking blood bank” donor. Although under specific conditions, a single individual may very well benefit from this source, the risk of transmitting a disease is high, even when donors are used that have been tested recently. Most expeditionary missions are land-based operations in third world countries. This means that infections like malaria and Chagas’ disease, even in previously healthy personnel, can and will be transmitted inadvertently from the donor pool to patients. Moreover, since blood is needed at unpredictable moments, the listed donors may not be available at the right moment for lots of reasons.29 The main reason to use fresh whole blood in theater is to treat major bleeding, due to lack of platelets in the patient, since liquid stored platelets are not available. In order to abandon the “walking blood bank”, an alternative for fresh whole blood was required.

DMSO frozen platelets have been transfused successfully since the 1970s.19,20 Despite the fact that their functional recovery is less than that of fresh liquid platelets (around 50%), the frozen platelets are more effective in stopping non-surgical bleeding compared to three day liquid stored platelets.21 Frozen platelets also show a higher capacity to bind factor V30 and a higher thromboxane A2 production after ADP stimulation.31 In addition, baboon frozen platelets have a higher “in vivo” survival and a stronger correction of aspirin induced prolonged bleeding time, compared to 5 day liquid stored platelets.32 With or without leukodepletion, platelets frozen with 4-6% DMSO may be stored at -80°C for at least 2 years.11 The possibility of adding frozen platelets to our line of products was therefore very logical to investigate.

Washing became part of the original thawing procedure of frozen platelets to reduce all possible adverse side effects of DMSO. When we visited Dr. Valeri


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to learn the freezing and thawing procedure of -80°C frozen platelets, he had modified the procedure in that postthaw washing was no longer required, because supernatant plasma, containing the majority of the DMSO, was removed prior to freezing. This modified procedure saves valuable time (±1 h), technical training and equipment (centrifuge) in theater. Since this was a major improvement (time-saving is life-saving), we used this modified freezing protocol.

Whereas Dr. Valeri uses saline, we use AB plasma to suspend the platelets after thaw, reasoning that a trauma patient needing red cells and platelets, almost certainly will also need plasma. Autologous plasma has been used previously for the suspension of thawed washed platelets.20 We chose not to use this option, because when using AB plasma and O platelets, the product can be used in every patient regardless of the ABO blood type. In our experience, the thawed platelets show clearly visible swirling in plasma and the product looks like a regular platelet unit obtained by apheresis. We found an “in vitro” recovery of ±77%, similar to the “in vitro” recovery reported by others.11,17-19

By the end of 2001 we implemented the use of frozen platelets in Bosnia, and abandoned the “walking blood bank” concept. Within half a year, two patients were treated that required platelet transfusion. One elderly woman with gunshot wounds in the pelvic region and one young soldier with acute ITP (viral). Both patients experienced unstoppable bleeding, due to a low platelet concentration. Although the platelet count barely rose after transfusion, the bleeding of each patient stopped within 20 min after transfusion of one thawed platelet concentrate in AB plasma. We have thus experienced that frozen platelets can be life-saving and that the use of a walking blood bank can be abolished when this product is available in theater. From that time onward, frozen platelets and frozen blood bank facilities always have been part of the standard equipment of Dutch deployed military hospitals.

Fresh frozen plasma (FFP) is normally kept at temperatures of around -20°C to -30°C with a shelf life from 1 to 2 years. Lower temperatures of at least -65°C even permit a storage period up to 7 years.9,10 Apheresis, leukodepleted plasma is frozen by Sanquin, according to the guidelines, and meets the requirements for FFP. The units are still frozen upon arrival at the MBB, not folded and do not fit into our standard rigid cardboard boxes. Since we wanted to maintain uniformity in our packing methods and -80°C transportation and storage temperature, we repacked the plasma units.

Refrozen (-30°C), previously thawed FFP and even liquid stored plasma, have been shown to be safe and effective, although factor V and FVIII:C levels are significantly decreased with 10-30%.33-37 We observed a slight decrease in factor V and VIII levels due to the extra thaw-freeze cycle, however since we only have used AB plasma which has the advantage of high factor VIII levels to start with, factor V and VIII levels are still well above the 0.7 U/mL. It is likely, but still unproven, that these refrozen DFP units can be stored for 7 years at -80°C as described for fresh frozen plasma at temperatures below -65°C.9,10

In literature it is suggested that breakage of PVC at -80°C is due to freezing and cannot be prevented, instead new plastics are studied to avoid breakage.38 Strong reduction of PVC breakage by means of folding the blood bag to protect vulnerable parts and by putting it in an overwrap bag and a rigid cardboard box has been described by Valeri et al.11,39 The overwrap bag prevents contact of the product bag with the cardboard box which reduces breakage and is also used during thaw to protect the water bath against contamination, in case the product bag is broken. Using the same method in combination with a vacuum-sealed overwrap bag, we also could strongly reduce breakage of standard PVC bags stored at -80°C; breakage was reduced to 0.0%, even after transportation on “dry ice”.

To date, the (inter)national guidelines advocate the use of frozen cellular blood products only for rare blood types and patients with multiple alloantibodies. We have shown that the frozen blood bank concept can be effectively used in a military environment and are convinced that it can also be very useful in the civilian community. A frozen blood bank facility with a stock of frozen universal donor products can effectively be used in remote areas, to compensate for periods when no donors are available (holiday periods) and when suddenly many patients are in need for blood products. In addition, a quarantine period similar to that currently used for plasma can also be used for frozen cellular blood products to reduce risks of transmitting diseases.


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Conclusions

The quality of our stock of frozen blood products is in compliance with the European and US standards.9,10 The products can easily, effectively and safely be used in deployed blood bank units. Deployed military hospitals become thus less dependent on their home countries or on a “walking blood bank”. Products can be produced on demand and blood “spillage” due to expiration can be reduced to the absolute minimum. The logistic burden can be reduced since less or no shipments are required to support for the blood product inventory abroad.

Acknowledgments

We like to thank Dr. C.R. Valeri for providing instructions, protocols, advice and technical support, Sanquin Blood Bank Southwest for providing liquid blood products and leukodepletion data. Furthermore we thank the pharmacists S.M.G.E. van Grinsven, J. Chin and H.M. Meinen, for their leading role in operating the deployed blood bank in Iraq and S.F. Bus, M. Hakvoort, D. van Zandwijk and R. Nieuwenhuizen for their technical assistance. The Netherlands Ministry of Defense financially supported this study. The opinions or assertions contained herein are those of the authors and are not to be construed as official or reflecting the views of the Ministry of Defense.

References

1. Champion HR, Bellamy RF, Roberts CP, Leppaniemi A. A profile of combat injury. J.Trauma 2003;54(5 Suppl):S13-S19.

2. Armand R, Hess JR. Treating coagulopathy in trauma patients. Transfus.Med Rev. 2003;17(3):223-31.

3. Hardy JF, de Moerloose P, Samama CM. The coagulopathy of massive transfusion. Vox Sang. 2005;89(3):123-7.

4. Scott KL, Lecak J, Acker JP. Biopreservation of red blood cells: past, present, and future. Transfus.Med Rev. 2005;19(2):127-42.

5. Blajchman MA. Novel platelet products, substitutes and alternatives. Transfus.Clin.Biol. 2001;8(3):267-71.

6. Buehler PW, Alayash AI. Toxicities of hemoglobin solutions: in search of in-vitro and in-vivo model systems. Transfusion 2004;44(10):1516-30.

7. Valeri CR, Pivacek LE, Gray AD, Cassidy GP, Leavy ME, Dennis RC, Melaragno AJ, Niehoff J, Yeston N, Emerson CP. The safety and therapeutic effectiveness of human red cells stored at -80 degrees C for as long as 21 years. Transfusion 1989;29(5):429-37.

8. Valeri CR, Srey R, Tilahun D, Ragno G. The in vitro quality of red blood cells frozen with 40 percent (wt/vol) glycerol at -80 degrees C for 14 years, deglycerolized with the Haemonetics ACP 215, and stored at 4 degrees C in additive solution-1 or additive solution-3 for up to 3 weeks. Transfusion 2004;44(7):990-5.

9. Standards for Blood Banks and Transfusion Services. 23rd edition. 2004. Bethesda, AABB.

10. Guide to the Preparation, Use and Quality Assurance of Blood Components. 11th edition. 2005. Strasbourg, Council of Europe.

11. Valeri CR, Srey R, Lane JP, Ragno G. Effect of WBC reduction and storage temperature on PLTs frozen with 6 percent DMSO for as long as 3 years. Transfusion 2003;43(8):1162-7.

12. Sowemimo-Coker SO. Red blood cell hemolysis during processing. Transfus.Med.Rev. 2002;16(1):46-60.

13. Mollison PL. Blood Transfusion in Clinical Medicine.10th edition, 1997,p 302. Blackwell Science.




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16. Bandarenko N, Hay SN, Holmberg J, Whitley P, Taylor HL, Moroff G, Rose L, Kowalsky R, Brumit M, Rose M, et al. Extended storage of AS-1 and AS-3 leukoreduced red blood cells for 15 days after deglycerolization and resuspension in AS-3 using an automated closed system. Transfusion 2004;44(11):1656-62.

17. Melaragno AJ, Abdu WA, Katchis RJ, Vecchione JJ, Valeri CR. Cryopreservation of platelets isolated with the IBM 2997 blood cell separator: a rapid and simplified approach. Vox Sang. 1982;43(6):321-6.

18. Vecchione JJ, Chomicz SM, Emerson CP, Valeri CR. Cryopreservation of human platelets isolated by discontinuous-flow centrifugation using the Haemonetics Model 30 Blood Processor. Transfusion 1980;20(4):393-400.

19. Schiffer CA, Aisner J, Wiernik PH. Clinical experience with transfusion of cryopreserved platelets. Br.J.Haematol. 1976;34(3):377-85.

20. Schiffer CA, Aisner J, Dutcher JP, Daly PA, Wiernik PH. A clinical program of platelet cryopreservation. Prog.Clin.Biol.Res. 1982;88:165-80.

21. Khuri SF, Healey N, MacGregor H, Barnard MR, Szymanski IO, Birjiniuk V, Michelson AD, Gagnon DR, Valeri CR. Comparison of the effects of transfusions of cryopreserved and liquid-preserved platelets on hemostasis and blood loss after cardiopulmonary bypass. J.Thorac.Cardiovasc.Surg. 1999;117(1):172-83.

22. Lelkens CCM, Noorman F, Koning JG, Truijens-de Lange R, Stekkinger PS, Bakker JC, Lagerberg JWM, Brand A, Verhoeven AJ. Stability after thawing of RBCs frozen with the high- and low-glycerol method. Transfusion 2003;43(2):157-64.

23. Valeri CR, Sims KL, Bates JF, Reichman D, Lindberg JR, Wilson AC. An integrated liquid-frozen blood banking system. Vox Sang. 1983;45(1):25-39.


25. Valeri CR, Brodine CE, Moss GE. Use of frozen blood in Vietnam. Bibl.Haematol. 1968;29:735-8.

26. Rosenblatt MS, Hirsch EF, Valeri CR. Frozen red blood cells in combat casualty care: clinical and logistical considerations. Mil.Med. 1994;159(5):392-7.

27. Cinat ME, Wallace WC, Nastanski F, West J, Sloan S, Ocariz J, Wilson SE. Improved survival following massive transfusion in patients who have undergone trauma. Arch.Surg. 1999;134(9):964-8.

28. Ho AMH, Karmakar MK, Dion PW. Are we giving enough coagulation factors during major trauma resuscitation? Am.J.Surg. 2005;190(3):479-84.

29. Reade MC. Blood products on operational deployments. ADF Health 2001;2:65-70.

30. Barnard MR, MacGregor H, Ragno G, Pivacek LE, Khuri SF, Michelson AD, Valeri CR. Fresh, liquid-preserved, and cryopreserved platelets: adhesive surface receptors and membrane procoagulant activity. Transfusion 1999;39(8):880-8.

31. Valeri CR, MacGregor H, Ragno G. Correlation between in vitro aggregation and thromboxane A2 production in fresh, liquid-preserved, and cryopreserved human platelets: effect of agonists, pH, and plasma and saline resuspension. Transfusion 2005;45(4):596-603.

32. Valeri CR, MacGregor H, Giorgio A, Ragno G. Circulation and hemostatic function of autologous fresh, liquid-preserved, and cryopreserved baboon platelets transfused to correct an aspirin-induced thrombocytopathy. Transfusion 2002;42(9):1206-16.

33. Milam JD, Buzzurro CJ, Austin SF, Stansberry SW. Stability of factors V and VIII in thawed fresh frozen plasma units. Transfusion 1980;20(5):546-8.

34. Smak Gregoor PJ, Harvey MS, Briet E, Brand A. Coagulation parameters of CPD fresh-frozen plasma and CPD cryoprecipitate-poor plasma after storage at 4 degrees C for 28 days. Transfusion 1993;33(9):735-8.

35. Dzik WH, Riibner MA, Linehan SK. Refreezing previously thawed fresh-frozen plasma. Stability of coagulation factors V and VIII:C. Transfusion 1989;29(7):600-4.

36. Ben-Tal O, Zwang E, Eichel R, Badalbev T, Hareuveni M. Vitamin K-dependent coagulation factors and fibrinogen levels in FFP remain stable upon repeated freezing and thawing. Transfusion 2003;43(7):873-7.

37. Cardigan R, Lawrie AS, Mackie IJ, Williamson LM. The quality of fresh-frozen plasma produced from whole blood stored at 4 degrees C overnight. Transfusion 2005;45(8):1342-8.

38. Hmel PJ, Kennedy A, Quiles JG, Gorogias M, Seelbaugh JP, Morrissette CR, Van Ness K, Reid TJ. Physical and thermal properties of blood storage bags: implications for shipping frozen components on dry ice. Transfusion 2002;42(7):836-46.

39. Valeri CR, Ragno G. Breakage rate for red blood cells frozen with 40 percent (wt/vol) glycerol in 800-mL polyvinylchloride plastic bags stored in rigid cardboard boxes at -80 degrees C. Transfusion 2005;45(5):822-3.

Susan J. Neuhaus1, Ken Wishaw2 and Charles C.M. Lelkens3

1Department of Surgery, University of Adelaide and Royal Adelaide Hospital, SA, Australia2Nambour General Hospital, Nambour, QLD, Australia

3Military Blood Bank, Leiden, the Netherlands

Med J Aust. 2010 Feb 15; 192(4):203-5

Chapter

Australian experience with frozen blood products on military operations

4

Chapter 4 Australian experience with frozen blood products on military operations

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Abstract

Historically, the Australian Defence Force (ADF) has sourced all its blood supplies from the Australian Red Cross Blood Service. Recent ADF operations in the Middle East have highlighted a need to rely on other nations’ blood supply systems.

In 2008, the ADF embedded a surgical and intensive care team into the Netherlands-led forward health facility at the Uruzgan Medical Centre (UMC) at Tarin Kowt in Afghanistan. To date, three teams have provided 2-month rotations as part of the North Atlantic Treaty Organization International Security Assistance Force in Afghanistan.

The Netherlands armed forces use a sophisticated system for supply of liquid and frozen blood products (frozen red cells, plasma and platelets).

We review Australian experience with the Dutch system of supplying blood products for major trauma resuscitation in Afghanistan.

Introduction

Exsanguinating haemorrhage is the major cause of death in current military conflict operations.1,2 Recent data from Afghanistan and Iraq have shown that up to 15% of combat casualties require massive transfusion for traumatic injuries, with a mortality rate of 20% -50%.3,4 This is a significantly higher proportion than in most civilian trauma centres, reflecting the severity and polytrauma of blast-associated injuries.

Current battlefield resuscitation practice focuses on early diagnosis and management of haemorrhagic shock, with the aim of rapidly reversing the lethal triad of acidosis, hypothermia, and coagulopathy, using aggressive resuscitation with a 1:1:1 ratio of red cells to plasma to platelets, and recombinant factor VIIa if required.5,6 Surgical intervention is focused on controlling haemorrhage and contamination (damage control), with definitive care delayed until normal coagulation and metabolic function have been restored.

Current options for provision of blood and blood products include fresh liquid supply, “walking donor panels” (people who are prepared to be called on to meet a particular emergency), and frozen blood products.

Supply of fresh liquid blood to combat areas is logistically challenging. Some military forces use pre-screened “walking donor panels”. However, this practice carries a risk of disease transmission, relies on the availability of appropriate donors, and restricts their duties for a period after donation, with implications for combat readiness (e.g., infantry soldiers are probably not fit for combat for at least 24 hours). Blood salvage is not used in the combat environment.7

Historically, the Australian Defence Force (ADF) has relied on fresh blood supplies from the Australian Red Cross Blood Service. But in recent ADF operations in the Middle East, where obtaining supplies of fresh blood is not feasible because of time and distance factors, Australia has been reliant on a Dutch national supply system. The Netherlands armed forces use a sophisticated system for supply of liquid and frozen blood products (red cells, plasma and platelets).8 Here, we review the ADF’s experience with frozen blood products in Afghanistan.

The Netherlands frozen blood bankFrozen red cells have been used on deployed military operations since the Vietnam War.9 Using frozen blood during military operations is appealing, as


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it avoids many logistic resupply issues and extends the shelf life compared with liquid blood. Deep-frozen red cells can be stored at -80°C for at least 10 years,10 deep-frozen plasma (DFP) for 7 years and deep-frozen platelets for 2 years.11

The Netherlands developed a state-of-the-art frozen blood component system during North Atlantic Treaty Organization operations in Bosnia in the 1990s.7 In 2002, universal frozen blood products were introduced, including a frozen platelet system. This is the system now in place at the Uruzgan Medical Centre (UMC) at Tarin Kowt in Afghanistan, where the ADF embedded a surgical and intensive care team into the Netherlands-led forward health facility in 2008.

All blood products are sourced from the Netherlands’ Sanquin Blood Supply Foundation and derived from an unpaid volunteer donor pool.8 Blood is screened and tested in accordance with national and international guidelines. To minimise the risk of transmission of Creutzfeldt - Jakob disease, blood donations are not accepted from people who have a history of transfusion or have lived in the United Kingdom for more than 6 months between 1990 and 1996.12,13

The Netherlands Military Blood Bank is supplied with universal donor red cells and platelets (group O Rh D-positive and -negative), as well as universal donor plasma (group AB). UMC routinely holds 30 units of thawed, liquid red cells, 150 units of frozen red cells, 60-70 units of DFP and 40 units of frozen platelets, with increased demand met by resupply from the Netherlands.

Red blood cellsRed cells are frozen in 40% glycerol (w/v) and transported from the Netherlands in temperature-monitored containers (TempTale® [Sensitech, Beverly, Mass, USA]) maintained at less than -65°C, and then transferred to a -80°C freezer at UMC.(See picture Above on page 74)

Cells are processed in a fully closed, semi-automated ACP 215 processor (Haemonetics, Braintree, Mass, USA). They are deglycerolised with 12% sodium chloride, washed with 0.9% sodium chloride and 0.2% glucose, and resuspended in a citrate-containing nutritional solution (AS-3 Nutricel [Gambro BCT, Lakewood, CO, USA]).(See picture Left on page 74)

Three units of red cells can be produced every 90 minutes (this includes 30 min thawing time). Regular processing allows liquid blood stores to be replenished to maintain an immediately available supply. The shelf life of red cells after thawing and processing is 14 days at 2-6°C. Cell vitality studies show a mean freeze-thaw-wash recovery value of 90%, a mean 24 hour posttransfusion

survival rate of 85%, normal or slightly impaired oxygen transport function, and minimal haemolysis.10

PlasmaGroup AB Rh (D) positive plasma is provided as DFP using single-donor apheresis. Plasma is citrated and leukocyte depleted. DFP is transported at a temperature of less than -65°C and stored at -80°C. Thawing takes about 30 minutes. The performance of DFP is almost identical to that of fresh frozen plasma.9

PlateletsLeukocyte-depleted, volume-reduced frozen platelets are obtained by apheresis from a single donor, cryoprotected in a solution of 4-6% dimethyl sulfoxide and stored at -80°C.14 Each unit contains about 300 x 109 platelets (equivalent to 5-6 donor units of fresh buffy coat platelets). The freezing process induces both morphological and functional changes.15,16 Compared with liquid stored platelets, frozen platelets demonstrate a higher capacity to bind factor V and higher thromboxane A2 production after stimulation with adenosine diphosphate.16 The thawing time of platelets is about 5 minutes. They are suspended in one unit of thawed DFP. Processing with dimethyl sulfoxide gives frozen platelets a uniquely pungent odour.

The Australian experienceSince ADF medical officers joined UMC in 2008, three teams have provided 2-month rotations as part of the International Security Assistance Force in Afghanistan. A prospective database was maintained during the ADF rotations. Of 158 patients undergoing surgery by Australian surgical teams, 17 received blood products intraoperatively (132 red cell units, 75 DFP units, 22 platelet units). One patient received recombinant factor VIIa. The predominant indication for surgery was blast- or gunshot-related injury. Over 90% of patients were Afghan nationals. No Australian service personnel received blood products during these periods. The following case studies illustrate some issues related to the use of frozen blood products.

Case study 1On 2 February 2009 at 08:20, a suicide bomber self-detonated at the Afghan


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National Police training barracks in Tarin Kowt, resulting in 22 deaths. A 25-year-old man presented with a blood pressure of 115/80 mm Hg, a pulse of 120 beats per min, and a Glasgow Coma Scale score of 15. He had a left flank entry wound with no exit wound. His abdomen was distended, and frank haematuria was apparent in urine collected via a urinary catheter. A Focused Assessment with Sonography for Trauma (FAST) scan gave positive results. There was no obvious chest injury. His initial haemoglobin level was 154 g/L. Resuscitation included rapid infusion of 5 units of non-crossmatched red cells before surgery. After induction of anaesthesia, the patient received 1L normal saline, and 30 minutes later, four units of crossmatched red cells, two units of DFP and two units of DFP with platelets. Laparotomy revealed free intraperitoneal bleeding, jejunal blast perforations, liver laceration and non-expanding left pelvic haematoma. The spleen was intact. There was a colonic contusion and a mesocolic haematoma. The lower pole of the kidney was macerated, with bleeding into the renal pedicle. A damage control approach was adopted, involving stapled exclusion of the small bowel, left nephrectomy, packing and laparotomy. The patient was transferred to the intensive care unit (ICU) for a planned return to theatre within 24 hours. The patient remained haemodynamically stable in the ICU. Coagulation studies the following morning showed an activated partial thromboplastin time (APTT) of 14.6s (reference range [RR] 28-40s); and a prothrombin time (PT) of 19.9s (RR 10-16s). At definitive laparotomy, surgical packs were densely adherent, but there was no evidence of ongoing bleeding. The patient’s further postoperative course was largely unremarkable. On discharge from the ICU at day 3, his haemoglobin level was 114 g/L.

Case study 2On 28 February 2009 at 14:50, a 20-year-old man presented with gunshot wounds to his left foot and upper right thigh. There were no other injuries. On arrival, he was severely shocked, with poor capillary perfusion, systolic blood pressure of 60 mm Hg and a pulse of 130 beats per min.

He was distressed and, within minutes, lost consciousness. Examination revealed anterior entry and posterior exit wounds to the right thigh, with a large haematoma and compartment syndrome.

At 15:10, he was in the operating room. His core temperature was 33.5°C, and blood gas measurements showed severe acidosis, with a base excess of - 21mmol/L. Operative findings were laceration of the superficial femoral artery,

superficial femoral vein and profunda femoris vessels, with extensive cavitation, soft tissue damage and multiple comminuted bone fragments. The patient was clinically coagulopathic. A temporary vascular shunt was inserted and fasciotomies were performed, with debridement and external fixation. Bleeding from the posterior thigh exit wound was controlled with HemCon® bandage (HemCon Medical Technologies, Portland, OR, USA) and pressure. Initial coagulation studies at 16:30 showed an APTT of 92s (RR 28-40s) and a PT of 19.6s (RR 10-16s). There was significant oozing from all operative sites.

After receiving 1 L of saline in the emergency room, the patient was given four non-crossmatched units of red cells, three units of DFP and one unit of DFP with platelets within 40 minutes. A further eight units of red cells, six units of DFP and two units of DFP with platelets were administered over the following 3 hours. Active warming was used throughout surgery. After leg reperfusion, the patient required intravenous administration of calcium gluconate, insulin and dextrose because there were electrocardiographic signs of severe hyperkalaemia.

Two doses of recombinant factor VIIa were given (initially 100 μg/kg and then another 60 μg/kg midway through the 4-hour procedure). At 20:15, the patient’s temperature was 37.4 °C, base excess was −1mmol/L, and coagulation times were essentially normal (APTT 46s; PT 13.8s). There were no clinical signs of coagulopathy by 22:30.

Resuscitation continued immediately after the operation, with the patient receiving a total of 16 units of red cells, 15 units of DFP and four units of DFP with platelets. In anticipation of potential myoglobinaemic renal failure, a forced diuresis regimen was undertaken for the next 24 hours. The patient’s condition remained stable, without signs of clinical or laboratory coagulopathy. He was evacuated to another facility 24 hours after surgery.

Lessons learnedThese cases demonstrate the use of integrated liquid and frozen blood components for patients with battlefield trauma. Non-crossmatched units were used immediately, with formal crossmatching, typing, antibody screening and agglutination tests performed as soon as practicable. The cases illustrate that, in an austere environment or during mass casualty events, decisions must frequently be made on clinical grounds, often without recourse to investigations.17 Except in Case study 2 (described here), there was no clinical evidence of coagulopathy in patients treated with blood products at UMC - an unusual observation given the


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severity of the injuries (median injury severity score, 41; range 4-75). Aggressive management of hypothermia was practised, including maintaining emergency and operating room temperatures above 28°C, active patient warming, and warming of all intravenous and irrigation fluids to 40°C. Our experience with “re-look” laparotomy was that packs were densely adherent. Despite simple measures such as irrigation to facilitate removal of packs, serosal damage and liver surface bleeding occurred, but these were easily controlled. We postulate this may be a thrombin-type effect related to activated platelets. This effect has not previously been reported and requires investigation. Recent operations in the Middle East have identified potential requirements for frozen blood products, particularly platelets, in military trauma settings. In partnership with the Australian Red Cross Blood Service, the ADF is investigating the utility of frozen platelets within the Australian therapeutic regulatory framework. Australian doctors should become familiar with the use and efficacy of these blood components.

Conclusions

Integrated fresh-frozen blood banking provides flexible and efficient use of blood products in a military setting. Despite infrastructure costs, it is logistically appealing and minimises wastage.

As the pendulum swings towards early component therapy in trauma resuscitation, the use of an integrated fresh - frozen blood bank may also help to meet the logistical and geographical challenges of supplying blood products to people in rural Australia.

Acknowledgements

We thank Colonel Leonard Brennan of the ADF, and Lieutenant Colonel Willem Sandberg, Warrant Officer Michael Lam, Corporal Freek Dekker and Sergeant Jorrit ten Broeke of the Royal Netherlands Army for their valuable assistance, advice and patience. The opinions or assertions contained in this article are those of the authors and are not to be construed as official or reflecting the views of the ADF.

Above:Inside the blood bank container

Left:Deglycerolising red blood cells in the ACP 215 processor


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Competing interests

None identified.

References

1. Hutt J, Wallis L. Blood products in trauma resuscitation. J.R.Army Med Corps 2006;152(3):121-7.

2. Alam HB, Rhee P. New developments in fluid resuscitation. Surg.Clin.North Am. 2007;87(1):55-72.

3. Borgman MA, Spinella PC, Perkins JG, Grathwohl KW, Repine T, Beekley AC, Sebesta J, Jenkins D, Wade CE, Holcomb JB. The ratio of blood products transfused affects mortality in patients receiving massive transfusions at a combat support hospital. J.Trauma 2007;63(4):805-13.

4. Huber-Wagner S, Qvick M, Mussack T, Euler E, Kay MV, Mutschler W, Kanz KG. Massive blood transfusion and outcome in 1062 polytrauma patients: a prospective study based on the Trauma Registry of the German Trauma Society. Vox Sang. 2007;92(1):69-78.

5. Holcomb JB, Wade CE, Michalek JE, Chisholm GB, Zarzabal LA, Schreiber MA, Gonzalez EA, Pomper GJ, Perkins JG, Spinella PC, et al. Increased plasma and platelet to red blood cell ratios improves outcome in 466 massively transfused civilian trauma patients. Ann.Surg. 2008;248(3):447-58.

6. Defense Medical Readiness Training Institute. Joint theater trauma system clinical practice guideline, 25 October 2007. http://www.dmrti.army.mil/documents (accessed Mar 2009).

7. Reade MC. Blood products on operational deployments. ADF Health 2001;2:65-70.



10. Valeri CR, Pivacek LE, Gray AD, Cassidy GP, Leavy ME, Dennis RC, Melaragno AJ, Niehoff J, Yeston N, Emerson CP. The safety and therapeutic effectiveness of human red cells stored at -80 degrees C for as long as 21 years. Transfusion 1989;29(5):429-37.


12. Van Everdingen JJE, Klazinga NS, Casparie AF. Blood transfusion policy in Dutch hospitals. 1988. Int.J.Health Care Qual.Assur. 2007;20(1):77-83.


14. Valeri CR, Ragno G, Khuri S. Freezing human platelets with 6 percent dimethyl sulfoxide with removal of the supernatant solution before freezing and storage at -80 degrees C without postthaw processing. Transfusion 2005;45(12):1890-8.

Chapter 4

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Prolonged postthaw shelf life of red cells frozen without prefreeze removal of excess glycerol

515. Barnard MR, MacGregor H, Ragno G, Pivacek LE, Khuri SF, Michelson AD, Valeri CR. Fresh, liquid-preserved, and cryopreserved platelets: adhesive surface receptors and membrane procoagulant activity. Transfusion 1999;39(8):880-8.


17. Neuhaus SJ, Sharwood PF, Rosenfeld JV. Terrorism and blast explosions: lessons for the Australian surgical community. ANZ.J.Surg. 2006;76(7):637-44.

Charles C.M. Lelkens1, Dirk de Korte2 and Johan W.M. Lagerberg2

1Royal Netherlands Navy, Medical Corps, Retired, Kortgene, the Netherlands2Blood Cell Research, Sanquin Research, Amsterdam, the Netherlands

Vox Sang. 2015;108(3):219-25.

Chapter 5 Prolonged postthaw shelf life of red cells frozen without prefreeze removal of excess glycerol

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Abstract

Background and Objectives. The use of a functionally closed system (ACP215, Haemonetics) for (de)glycerolization of RBCs allows for prolonged postthaw storage. Currently, glycerolization is followed by supernatant glycerol reduction before freezing. The aim of this study was to investigate the influence of supernatant glycerol reduction before freezing on the stability of thawed, deglycerolized RBCs during subsequent cold storage.

Materials and Methods. Leukoreduced RBCs were stored for 6 days at 2-6°C before glycerolization. The RBCs were pooled and split, and glycerol was added using the ACP215 to a final concentration of 40%. Units were either frozen as such (n = 4) or supernatant reduced before freezing (n = 4). After storage at -80°C, the units were thawed, deglycerolized and resuspended in SAGM. An additional sixteen units, frozen without supernatant reduction, were resuspended in either AS-3 (n = 8) or SAGM (n = 8) after deglycerolization. During cold storage (2-6°C), the red cells were analyzed for their stability and in vitro quality.

Results. The freeze-thaw-wash recovery was comparable for both volume reduced and non-reduced units. During postthaw storage in SAGM, non-glycerol reduced units showed significantly less potassium leakage and hemolysis and higher ATP levels. AS-3 strongly reduced hemolysis during postthaw storage of non-glycerol reduced units: hemolysis remained below 0.8% for up to 28 days of storage.

Conclusion. Omitting glycerol supernatant reduction before freezing simplifies the cryopreservation procedure and increases the stability and therefore the outdating period of thawed RBCs. This increases the practical applicability of cryopreserved RBCs in both civil (rare blood) and military blood transfusion practice.

Introduction

Meeting transfusion needs under military or other austere conditions is a logistic challenge of considerable proportions. Usually blood products have to be transported over large distances and once in theater, inventory management is hampered by unpredictable needs at unpredictable times. Moreover, in spite of technological developments over the past decades, massive blood loss remains a major cause of death on the battlefield.1-6

The standard shelf lives of essential blood components like red cells and platelets are relatively short, thus easily leading to either shortages or wastage. In civilian life, this also applies to managing an inventory of rare blood groups for high frequency antigen negative patients in need of a blood transfusion.

One method of extending these shelf lives from days and weeks to at least some years is cryopreservation. With regard to red cells, this requires the use of a cryoprotectant like glycerol in either low (around 20%)7,8 or high (around 40%) concentration.9-11

Of these methods, the high-glycerol method (HGM) is now mainly used, because mechanical freezers can be used to store the frozen products at -80°C or below, in contrast with the low-glycerol method that is dependent on the use of liquid nitrogen (≤ -140°C).

Initially, the actual use of frozen RBCs was hampered mainly by processing time and a 24 h outdating period, due to potential bacterial contamination if thawing and washing is performed in a non-closed system.12,13

The introduction of an automated cell processing system (ACP215, Haemonetics Braintree, MA) meant a major step forward. With this system, it became possible to both glycerolize and deglycerolize RBCs in a functionally closed system, resulting in reduced potential for bacterial contamination and allowing prolonged postthaw storage.14,15 Using the ACP215, all fluids needed for the glycerolization and deglycerolization procedure are administered through 0.22 μm bacteria barrier filters, and all tubing connections are achieved using a sterile connection device to maintain a closed system. Looking at hemolysis in the unit to be transfused as a pretransfusion criterion, the shelf life of the thawed and washed red cell product became largely dependent upon the final storage solution. Thus, for SAGM the shelf life at 2-6°C was limited to 48 to 72 h,16 whereas the use of AS-3 enabled a postthaw shelf life of at least 14 days, while retaining acceptable RBC quality.15


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One of the most commonly used freezing methods based on a high-glycerol concentration is described by Valeri.11 After the addition of the glycerol to the appropriate concentration, the volume of the glycerolized unit is reduced and the hematocrit of the suspension is increased by removing supernatant glycerol in a centrifugation step prior to freezing. This centrifugation step is not only laborious, but also induces a risk of breakage of the bag. In addition, centrifugation force might have a negative effect on the integrity of red cells.17,18

Interestingly, List et al.19 recently investigated a modified method of glycerolization and deglycerolization using the ACP215, which eliminates the centrifugation step that reduces glycerolized RBC supernatant. The deglycerolized RBCs complied with the CE and AABB standards. Although the study was not designed as a comparison with the conventional method, there were indications that skipping the removal of the supernatant glycerol before freezing, could result in a longer postthaw storage shelf life.

In the first part of this study, therefore, we compared both methods in a paired study. Two RBCs in SAGM were pooled and split and glycerolized using the ACP215. One of the units was glycerol reduced before freezing (prefreeze volume reduction, PFVR) while the other was frozen as such (no-PFVR). After thawing and deglycerolization, thawed RBCs were resuspended in SAGM and stored at 2-6 °C. In the second part of the study, we investigated the possibility of extending the postthaw shelf life even beyond 14 days, by using the no-PFVR method in combination with our previously described modified washing procedure (phosphate buffered saline instead of the 0.2% glucose/0.9% saline solution) and using AS-3 as the final storage solution).16


Blood collection and processingAll blood donors met standard donation criteria and gave their written informed consent in accordance to the institution’s guidelines and practices. The blood studies were approved by the institutional medical ethical committees in accordance with the standards laid down in the 1964 Declaration of Helsinki.

Whole blood (WB), 500 mL ± 2%, was collected in quadruple bag, bottom-and-top collection systems containing 70 mL of CPD (Fresenius Kabi, Emmer Compascuum, the Netherlands). The WB units were placed on butane-1,4-diol

cooling plates (Compocool, Fresenius Kabi) to allow their temperatures to adjust to 20 to 24°C.20 After overnight hold, the WB units were processed according to the routine buffy coat procedure as previously described.21 Briefly, after a hard spin, the WB units were separated in components, using an automated blood component separator (Compomat, Fresenius Hemo-Care). RBC was diluted with 110 mL SAGM and WBC reduced using the inline leukoreduction filter (BioR, Fresenius Kabi). The RBC units were stored at 2-6°C for 6 days prior to glycerolization and freezing.

GlycerolizationAt the end of the 6-day storage period, the RBC was centrifuged at 3200 x g for 5 min. The additive solution was removed to produce RBC units with an Hct of 75 ± 5%. The RBC units were sterilely connected (SCD Terumo) to a disposable glycerolization set (LN225, Haemonetics), and glycerolization was completed using the ACP215. A volume of 6.2 mol/L (57%) glycerol solution (6.2 M, S.A.L.F. S.p.A., Cenate Sotto, Bergamo, Italy) was added to the red cells to achieve a final glycerol concentration of 40% (w/ v). The glycerolized cells were transferred to a 1000 mL PVC bag (VSE 6002Z, MacoPharma, Mouvaux, France) and, where indicated, the supernatant glycerol was removed after centrifugation at 1200 x g for 10 min to yield a glycerolized RBC unit with a Hct of 70 ± 5%. The RBC units were sealed in a plastic overwrap, placed in a cardboard box (Cekumed, Ooltgensplaat, the Netherlands) and frozen as described before.22 The frozen RBCs were stored at -80°C for at least 14 days.

Thawing, deglycerolization and storageThe frozen RBCs were thawed in a water bath maintained at 40°C, to achieve a surface temperature of 25-30°C as measured with a non-contact thermometer (Fluke 62 mini, Eindhoven, the Netherlands). The thawed RBC units were sterilely connected to a deglycerolization set (LN235, Haemonetics). We used the ACP215 deglycerolization protocol default settings for both PFVR and no-PFVR units. The thawed RBC units were washed in the ACP215 using 50 mL of 12% NaCl (Bio-Deglyc, Bioluz, Saint-Jean-de-Luz, France) followed by either 1.6 L 0.9% NaCl / 0.2% glucose (Bio-wash, Bioluz) for SAGM (Haemonetics) resuspension, or phosphate-buffered saline (PBS) for AS-3 (Haemonetics) resuspension. Upon completion of the deglycerolization process, the cells were washed with and finally resuspended in either SAGM or AS-3.


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The deglycerolized RBC units were stored at 2-6°C for up to 14 days during the first part of the study and up to 35 days during the second part. Supernatant osmolality of the deglycerolized units was measured using an Osmomat 030-D Cryoscopic osmometer (Gonotec, Berlin, Germany).

Hematological parameters and recoveryThe volume of the blood components was calculated from the net weight divided by the specific gravity: 1.027 g/mL for plasma, 1.100 for 40% glycerol and RBCs and 1.006 for SAGM. Using the hematocrit of the solution, the specific gravity of RBC in additive solution was calculated from these values.

Freeze-thaw-wash recovery was calculated by comparing the total Hb after deglycerolization to the total Hb prior to glycerolization. Total Hb was calculated as volume (L) x Hb concentration (g/L). Hb concentration and red blood cell count were determined on a hematology analyzer (Advia 2120, Siemens Medical Solutions Diagnostics, Breda, the Netherlands). Hct measurements were made with the spun capillary method. Hemolysis was determined as described previously.23 Briefly, cell-free supernatants were obtained by centrifugation of the RBC suspensions at 2000 x g for 10 min followed by an additional centrifugation of the supernatant at 12000 x g for 5 min. Free Hb was determined by absorbance measurement of supernatant at 415 or 514 nm by a spectrophotometer (Rosys Anthos ht3, Antos Labtec Instruments GmbH, Salzburg, Austria), with correction for plasma absorption if necessary. Hemolysis was expressed as a percentage of total Hb present in the RBC after correction for hematocrit.

Analysis of metabolic and cellular variablespH, Extracellular potassium, glucose and lactate were measured with a blood gas analyzer (Rapidlab 860, Siemens Medical Solutions Diagnostics).

ATP levels were determined in neutralized perchloric acid extracts. In short, extracts were made by diluting 600 μL of RBCs with 900 μL phosphate-buffered saline and then acidified with 60 μL of perchloric acid (70%wt/ vol). After 30 min on ice, extracts were centrifuged at 4°C at 6000 x g and 56 μL of 5 mol/L K2CO3 was added to 1 mL deproteinized supernatant for neutralization. Samples were kept frozen until analysis. Adenine nucleotides ATP, ADP and AMP were assayed using a HPLC method.24 Total adenylate was calculated as the sum of ATP, ADP and AMP levels.

Statistical analysisStatistical analyses were performed with standard software (Microsoft Excel, Bellevue, MA). Results are shown as mean ± SD with the number of observations given between parentheses. T-tests were used to determine the significance of differences between two groups. A p value of <0.05 was considered significant.

Results

In the first part of the study, pooled and split RBCs in SAGM were used. The volume and hemoglobin content of the units before glycerolization were comparable (see Table 1). As expected, units that were glycerolized without prefreeze volume reduction (no-PFVR) showed a significant larger volume and lower hematocrit as compared to units that were centrifuged before freezing (PFVR). Hemoglobin content of the frozen units was comparable for both groups. The volume of the washing fluid used in the deglycerolization process was larger for no-PFVR units than for PFVR units. The difference in volume can be completely accounted for by the lower hematocrit, and thus larger supernatant volume, of the no-PFVR units. The loss of hemoglobin in the washing fluid was higher for the no-PFVR units, resulting in deglycerolized products with slightly lower Hb content and hematocrit. Freeze-thaw-wash recovery was comparable for both PFVR and no-PFVR units. The osmolality of the supernatant of all deglycerolized products was below 420 mOsm/kg and comparable for both groups (see Table 1). All deglycerolized units therefore had a residual glycerol concentration of less than 1% (w/v),25 which indicates adequate removal,12 essential for safe transfusion.

Within the first days of postthaw storage in SAGM at 2-6°C, the PFVR units already show a significantly higher potassium leakage than the no-PFVR units (Fig. 1a). Also hemolysis during postthaw storage was significantly higher in the PFVR units (Fig. 1b). With the no-PFVR units, hemolysis remained below 0.8% until day 9 of storage, whereas the PFVR units already reached this level within 2 to 4 days of storage. During storage, ATP levels declined (Fig. 1c), with no large differences between both groups. Also glucose consumption, lactate production and pH were comparable for both groups (data not shown).

It is well known that AS-3 is a better storage solution for thawed red cells than SAGM. To investigate whether this also holds for no-PFVR units, 16


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units of red cells were frozen using the no-PFVR method. After thawing and deglycerolization, eight units were resuspended in SAGM and eight in AS-3. To compensate for the lower pH of AS-3, cells that were to be resuspended in AS-3 were washed with PBS instead of the usual 0.2% glucose/0.9% NaCl mixture as described before.16

During storage, supernatant potassium levels increased with no difference between SAGM and AS-3 (Fig. 2a). Storage in AS-3 resulted in significantly lower hemolysis. In agreement with the first part of the study, the SAGM units reached the cut-off value of 0.8% hemolysis after 9 days of storage (Fig. 2b). With AS-3, even after 35 days of storage, all units remained below this value (maximum hemolysis was 0.68%). ATP levels dropped considerably during postthaw storage, without any significant difference between SAGM and AS-3 units (Fig. 2c). The decline in ATP was accompanied by an increase of both ADP and AMP (not shown). The total adenylate content proved to be better preserved in AS-3. At day 35 of storage, total adenylate levels were still about 5.1 μmol/g Hb, being 75% of the value immediately after thawing (Fig. 2d).

Discussion

The FDA has already approved storage of thawed, washed red cells for 14 days in AS-3 at 2-6°C, if processed in a fully closed system.12 A further extension of this period would be most welcome to enable a better inventory management and at the same time help limiting unnecessary wastage and shortages, specifically under circumstances where demand and supply are hard to predict, that is battlefield conditions as part of combat casualty care and in civilian life for those patients who are dependent upon rare blood groups in case of a transfusion. Some authors14,26 have previously claimed 21 day postthaw storage being possible, but the results failed to meet the CE requirements regarding the upper limit of hemolysis, being 0.8%, in all units tested.13 In the study presented here, we tried to prolong the postthaw shelf life of red cells at 2-6°C by modifying the existing HGM procedure. The first part comprised a comparison of the original HGM method as described by Valeri11 and the modification of that method without prefreeze volume reduction as described by List et al.19 The results of the first part of our study confirm the suggestion from List et al. that skipping the centrifugation step before freezing leads to a better postthaw shelf life of

Figure 1. In vitro quality of deglycerolized cells during post-thaw storage in SAGM.

2 RBC units were pooled and split and glycerolized using the ACP215. One unit was frozen with (F) and one unit without (M) pre-freeze volume reduction. After thawing and deglycerolization, RBCs were resuspended in SAG-M, stored at 2-6°C and analyzed for in vitro quality parameters: A. Supernatant potassium, B. Hemolysis, C. ATP content. Results given are the mean ± SD of 4 units.*: Significantly different (p<0.05, paired Student’s t-test) from the values obtained with volume reduction.

1


2 RBC units were pooled and split and glycerolized using the ACP215. One unit

was frozen with (○) and one unit without (●) pre-freeze volume reduction. After

thawing and deglycerolization, RBCs were resuspended in SAGM, stored at 2-6°C

and analyzed for in vitro quality parameters: A. Supernatant potassium, B.

Hemolysis, C. ATP content. Results given are the mean ± SD of 4 units.

*: Significantly different (p<0.05, paired Student’s t-test) from the values obtained

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previously frozen red cells at 2-6°C. Based on a maximum allowed hemolysis of 0.8%, omitting the prefreeze centrifugation step, prolonged the postthaw shelf life of RBC in SAGM from 2-3 days to about 9 days. Potassium leakage and hemolysis in the modified method are reduced, indicating that the modified freeze-thaw-wash method results in less red cell damage than the original HGM method as proposed by Valeri. Although the exact reason for this difference remains to be elucidated, most probably, skipping the centrifugation step in the glycerolization procedure plays an important role in minimizing shear stress on the red cells,17,18 resulting in less damage, and thus better storable red cells. A number of studies have looked at the effect that hematocrit may have on the freezing response of RBCs.27,28 Reduced packing of RBCs within the unfrozen fraction might also explain part of the improvement in postthaw quality of the RBCs in the no-PFVR group. The metabolic characteristics of the previously frozen red cells appear to be unaffected by the change in the procedure, as indicated by the comparable glucose, lactate, pH and ATP levels. Potential negative effects of omitting the prefreeze glycerol removal could be the slightly higher loss of hemoglobin in the washing fluid, probably due to overloading of the centrifugation bowl, and the required larger storage capacity, due to the increased volume of the frozen units. A larger bowl may help to minimize the loss of hemoglobin during the washing procedure.

In the second part of this study, we compared SAGM and AS-3 as final storage solutions. It is well known that thawed RBC store better in AS-3, which was confirmed in the current study. Especially, the presence of citrate in AS-3 seems to play an important role in maintaining RBC integrity.16,29 Normal ATP concentrations have shown to be necessary to prevent calcium-induced membrane loss by microvesiculation30 and to maintain active transport of negatively charged phospholipids, especially phosphatidyl serine, from the outer to the inner leaflet of the RBC membrane. This mechanism prevents RBC clearance from the circulation by macrophages.31 In a previous study, we showed that RBC ATP-levels declined more rapidly during post-thaw storage in AS-3 as compared to SAGM. This was most likely due to the lower intracellular pH of cells in AS-3, resulting in decreased ATP synthesis32,33 and could be overcome by the use of PBS (pH 7.4) instead of the normally used, acidic, saline/glucose solution.16 Also in this study, PBS-washed RBCs resuspended in AS-3 maintained their ATP levels during postthaw storage to a comparable level as cells resuspended in SAGM (Fig. 2c).

Figure 2. In vitro quality of thawed, deglycerolized RBCs, frozen without volume reduction, during storage in SAGM or AS-3.

RBC units were glycerolized using the ACP215 and frozen without volume reduction. After thawing and deglycerolization, RBCs were resuspended in SAGM (F) or AS-3 (M), stored at 2-6°C and analyzed for in vitro quality parameters: A. Supernatant potassium, B. Hemolysis, C. ATP content, D. total adenylate content.Results given are the mean ± SD of 8 units.*: Significantly different (p<0.05, unpaired Student’s t-test) from the values obtained in SAGM

2

Figure 2. In vitro quality of thawed, deglycerolized RBCs, frozen without volume

reduction, during storage in SAGM or AS-3.

RBC units were glycerolized using the ACP215 and frozen without volume

reduction. After thawing and deglycerolization, RBCs were resuspended in SAGM

(○) or AS-3 (●), stored at 2-6°C and analyzed for in vitro quality parameters: A.

Supernatant potassium, B. Hemolysis, C. ATP content, D. total adenylate content.

Results given are the mean ± SD of 8 units.

*: Significantly different (p<0.05, unpaired Student’s t-test) from the values obtained

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Combining the three modifications in the cryopreservation method, thus skipping of prefreeze volume reduction, washing with PBS and the use of AS-3 as additive solution, makes it possible to maintain hemolysis below 0.8% for at least 35 days after thawing.

For “in vivo” recovery of red cells, total adenylate is of utmost importance. Högman et al.34 have shown a good correlation between total adenylate and in vivo recovery (r = 0.88). To achieve an “in vivo” 24 hour recovery of at least 75%, as required by US and CE guidelines,13,35 82% of the original total adenylate is required.34 Based on this last parameter, thawed red cells, if washed with PBS, can be stored for 4 weeks in AS-3.

In conclusion, omitting prefreeze volume reduction not only simplifies the procedure, but also reduces red cell damage induced by the cryopreservation procedure. Based on hemolysis and total adenylate content, thawed RBC resuspended in SAGM can be stored up to 9 days at 2-6°C. With PBS as washing solution and AS-3 as additive solution, the shelf life can be extended to 28 days.The practical implications of the results obtained in this study, we believe, are important. Although the simple fact remains that the use of frozen red cells comes at undeniably higher cost, a postthaw shelf life prolonged to 28 days would help solving logistic issues in civilian and military transfusion practice, without compromising the required safety and quality.

References

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2. Bellamy RF, Maningas PA, Vayer JS. Epidemiology of trauma: military experience. Ann.Emerg.Med 1986;15(12):1384-8.

3. Holcomb JB, McMullin NR, Pearse L, Caruso J, Wade CE, Oetjen-Gerdes L, Champion HR, Lawnick M, Farr W, Rodriguez S, et al. Causes of death in U.S. Special Operations Forces in the global war on terrorism: 2001-2004. Ann.Surg. 2007;245(6):986-91.

4. Eastridge BJ, Hardin M, Cantrell J, Oetjen-Gerdes L, Zubko T, Mallak C, Wade CE, Simmons J, Mace J, Mabry R, et al. Died of wounds on the battlefield: causation and implications for improving combat casualty care. J.Trauma 2011;71(1 Suppl):S4-S8.

5. Pannell D, Brisebois R, Talbot M, Trottier V, Clement J, Garraway N, McAlister V, Tien HC. Causes of death in Canadian Forces members deployed to Afghanistan and implications on tactical combat casualty care provision. J.Trauma 2011;71(5 Suppl 1):S401-S407.

6. Eastridge BJ, Mabry RL, Seguin P, Cantrell J, Tops T, Uribe P, Mallett O, Zubko T, Oetjen-Gerdes L, Rasmussen TE, et al. Death on the battlefield (2001-2011): implications for the future of combat casualty care. J.Trauma Acute.Care Surg. 2012;73(6 Suppl 5):S431-S437.






12. Standards for Blood Banks and Blood Transfusion Services. 28th edition. 2012. Bethesda, AABB.




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29. Besselink GAJ, Ebbing IG, Hilarius PM, de Korte D, Verhoeven AJ, Lagerberg JWM. Composition of the additive solution affects red blood cell integrity after photodynamic treatment. Vox Sang. 2003;85(3):183-9.

30. Kamp D, Sieberg T, Haest CW. Inhibition and stimulation of phospholipid scrambling activity. Consequences for lipid asymmetry, echinocytosis, and microvesiculation of erythrocytes. Biochemistry 2001;40(31):9438-46.

31. Verhoeven AJ, Hilarius PM, Dekkers DWC, Lagerberg JWM, de Korte D. Prolonged storage of red blood cells affects aminophospholipid translocase activity. Vox Sang. 2006;91(3):244-51.

32. Guppy M, Attwood PV, Hansen IA, Sabaratnam R, Frisina J, Whisson ME. pH, temperature and lactate production in human red blood cells: implications for blood storage and glycolytic control. Vox Sang. 1992;62(2):70-5.

33. Hess JR, Rugg N, Joines AD, Gormas JF, Pratt PG, Silberstein EB, Greenwalt TJ. Buffering and dilution in red blood cell storage. Transfusion 2006;46(1):50-4.


35. Technical Manual AABB; 2002.


16. Lagerberg JWM, Truijens-de Lange R, de Korte D, Verhoeven AJ. Altered processing of thawed red cells to improve the in vitro quality during postthaw storage at 4 degrees C. Transfusion 2007;47(12):2242-9.


18. Leitner GC, Dettke M, List J, Worel N, Weigel G, Fischer MB. Red blood units collected from bone marrow harvests after mononuclear cell selection qualify for autologous use. Vox Sang. 2010;98(3 Pt 1):e284-e289.

19. List J, Horvath M, Leitner GC, Weigel G. Cryopreservation of red blood cell units with a modified method of glycerolization and deglycerolization with the ACP 215 device complies with American and European requirements. Immunohematology. 2012;28(2):67-73.


21. van der Meer P, Pietersz R, Hinloopen B, Dekker W, Reesink H. Automated separation of whole blood in top and bottom bags into components using the Compomat G4. Vox Sang. 1999;76(2):90-9.


23. de Korte D, Kleine M, Korsten HGH, Verhoeven AJ. Prolonged maintenance of 2,3-diphosphoglycerate acid and adenosine triphosphate in red blood cells during storage. Transfusion 2008;48(6):1081-9.

24. de Korte D, Haverkort WA, van Gennip AH, Roos D. Nucleotide profiles of normal human blood cells determined by high-performance liquid chromatography. Anal.Biochem. 1985;147(1):197-209.

25. Clinical and Practical Aspects of the Use of Frozen Blood. AABB; 1977.23-36 p.26. Bohonek M, Petras M, Turek I, Urbanova J, Hradek T, Chmatal P, Staroprazska V,

Kostirova J, Horcickova D, Duchkova S, et al. Quality evaluation of frozen apheresis red blood cell storage with 21-day postthaw storage in additive solution 3 and saline-adenine-glucose-mannitol: biochemical and chromium-51 recovery measures. Transfusion 2010;50(5):1007-13.

27. Pegg DE. The effect of cell concentration on the recovery of human erythrocytes after freezing and thawing in the presence of glycerol. Cryobiology 1981;18(3):221-8.

28. Mazur P, Cole KW. Influence of cell concentration on the contribution of unfrozen fraction and salt concentration to the survival of slowly frozen human erythrocytes. Cryobiology 1985;22(6):509-36.

Charles C.M. Lelkens1, Johan W.M. Lagerberg1,2 and Dirk de Korte1,2

1 Sanquin Research, Department Blood Cell Research, Amsterdam, the Netherlands, and Landsteiner Laboratory, Academic Medical Center, University of Amsterdam,

Amsterdam, the Netherlands2 Sanquin Blood Bank, Department Product and Process Development,

Amsterdam, the Netherlands

Accepted for publication in Transfusion Feb 1 2017

Chapter

The effect of prefreeze rejuvenation on postthaw storage of red cells in AS-3 and SAGM

6

Chapter 6 The effect of prefreeze rejuvenation on postthaw storage of red cells in AS-3 and SAGM

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Abstract

Background: We investigated if improving the metabolic status of red cell concentrates (RCC) before freezing could extend the postthaw shelf life beyond 14 days, while still meeting the requirements for hemolysis (0.8%) and total adenylate (>82% of original values).

Study Design and Methods: At day 8 after collection, four leukoreduced RCC in SAGM were pooled, mixed and split (n=4). Of these RCC, two were rejuvenated in Rejuvesol. In addition, two leukoreduced RCC in PAGGGM were pooled, mixed and split at day 8 after collection (n=4). All RCC were glycerolized, frozen and stored for at least two weeks at 80°C. After thawing and deglycerolization, from each pair, one RCC was resuspended in SAGM and one in AS-3. During postthaw storage at 2-6°C for 35 days, all RCC were weekly sampled and analyzed for hematological, metabolic and morphological parameters.

Results: Both Rejuvesol and PAGGGM treatment showed increased ATP, total adenylate and 2,3-DPG levels compared to untreated RCC. Regardless of prefreeze Rejuvesol or PAGGGM treatment, postthaw hemolysis remained < 0.8% during 7 days in SAGM and during 35 days in AS−3. At day 35 of postthaw storage in AS-3, total adenylate of non-rejuvenated RCC had decreased to 72% of original values, whereas in prefreeze Rejuvesol and PAGGGM-treated RCC they still were at 101% and 98% respectively.

Conclusion: Based on maximum allowable hemolysis of 0.8% and total adenylate content of > 82% of the original value, thawed, prefreeze Rejuvesol- or PAGGGM-treated RCC can be stored for 35 days at 2-6ºC in AS-3.

Introduction

Freezing red cells enables preservation for a period of at least ten years.1 After the mandatory deglycerolization procedure such cells can be stored for another 14 days at 2-6°C in AS-3, if processed in a fully closed system.1-3 At these temperatures the cell’s metabolism is depressed but not stopped, so inevitably the detrimental effects of storage will start to appear at some point in time. During storage, red cells undergo a series of biochemical and morphologic changes, adversely affecting their primary role to take up, transport and deliver oxygen to tissues at the microcirculatory level. This complex of changes, particularly prominent after 14 days of storage and onward,4 has a strongly interdependent character and is commonly referred to and known as the ”storage lesion”.

The biochemical changes include a progressive loss of 2,3-DPG and ATP, as well as a buildup of lactic acid and oxidative damage to the red cell membrane’s lipids, proteins and carbohydrates.5 Also, an increased presence of phosphatidylserine is found on the outer leaflet of the red cell membrane.6

The morphologic changes comprise a transition from biconcave discs to echinocytes and finally to spheres,7 accompanied by a shedding of microvesicles.8

One of the consequences of the “storage lesion” is that damage to the membrane and loss of lipids from that membrane, including the formation of vesicles, lead to a less favorable surface-to-volume ratio.9 This has a negative effect on the red cell’s deformability, necessary to survive repeated passages through the narrowest capillaries with a diameter about half the one of a normal red cell.9 Thus, it will also reduce the red cell’s survival after transfusion.10

Biochemical modification, also called rejuvenation, is a method known to restore the red cell’s ATP and 2,3-DPG content to normal or even supranormal levels.11 It will also, at least partially, restore the shape change from spherocytes and echinocytes back to discs,12 improve oxygen delivery,13 as well as the membrane’s elasticity14 and reverse post-storage RBC adhesion to endothelial cells.15

The concept of rejuvenation, using solutions containing Pyruvate, Inosine, Phosphate and Adenine with or without glucose (PIGPA and PIPA respectively) has been developed and successfully used, mainly to prevent discarding of outdated red cells,16,17 but has also been advocated as being important in specific clinical situations, such as hemorrhagic shock11 and cardiopulmonary bypass patients e.g.18 Other than the costs and the pre-storage labor, the downside of


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this procedure is the fact that rejuvenated red cells need to be washed before transfusion, because of the potential toxicity of the metabolites of inosine and adenine in the rejuvenation solution.11 When applied to red cells meant to be frozen in glycerol, this downside however is relative, because these cells already need a mandatory postthaw washing procedure before transfusion to bring the residual glycerol concentration down to less than 1% (w/v),19 in order to avoid hemolysis in the patient’s circulation.

In an earlier study,20 we confirmed the findings of List et al.21 that omitting the prefreeze centrifugation step after glycerolization prolongs the postthaw shelf life of previously frozen red cells in SAGM, even considerably more if combined with the use of Phosphate Buffered Saline (PBS) as alternative washing solution and AS-3 as the final storage medium.22

The primary aim of the study presented here was to see if the combination of the previous modifications and the use of red cells with improved “in vitro” characteristics, specifically regarding ATP and 2,3-DPG content, prior to freezing with the High Glycerol Method (40%), could even further extend the postthaw shelf life of these cells.

For this purpose, prior to freezing, red cells were rejuvenated using a PIPA solution (Rejuvesol, Citra Labs, Braintree, MA) in one arm of the study (further referred to as pf-R, prefreeze Rejuvesol), whereas in the other arm red cells were prefreeze stored in the recently developed RBC additive solution PAGGGM, (further referred to as pf-PAGGGM, prefreeze PAGGGM), designed to better maintain metabolic status during storage (Table 1).23,24

In both arms of the study we compared the postthaw stability and quality after resuspension in AS-3 and SAGM.

Materials and Methods

Study designFour units of leukoreduced RBC in SAGM were stored at 2-6°C. At day eight after collection, the four units were pooled, mixed and split. Of these units, two were rejuvenated according to the manufacturer’s protocol. Within one hour after rejuvenation, all four units were glycerolized, frozen and stored for at least two weeks at -80°C. After thawing and deglycerolization, from each pair one unit was resuspended in SAGM and one in AS-3.

In addition, two units of leukoreduced RBC in PAGGGM were stored at 2-6°C. At day eight after collection, the units were pooled, mixed and split. Both units were subsequently glycerolized, frozen and stored for at least two weeks at -80°C. After thawing and deglycerolization, one unit was resuspended in SAGM and one in AS-3. This sequence was repeated three times over the following days for the leukoreduced SAGM- and PAGGGM units.

Thus, the postthaw study in one arm comprised eight units treated with Rejuvesol (originally stored in SAGM), four of which were resuspended in AS-3 and four in SAGM and eight non-rejuvenated (originally stored in SAGM) units, four of which were resuspended in AS-3 and four in SAGM. The other arm of the postthaw study comprised eight units (originally stored in PAGGGM), four of which were resuspended in AS-3 and four in SAGM. The design and the time frame of the study from collection to postthaw storage is depicted in Fig.1.

Blood collection and processingAll (non-remunerated) volunteer blood donors met standard donation criteria and gave their written, informed consent, in accordance with the institution’s

Table 1. Composition of Rejuvesol and Additive Solutions used

Ingredients (mmol/L)

Rejuvesol PAGGGM SAGM AS-3

NaCl - - 150 70Na2HPO4

103 8 - -NaH2PO4

52 8 - 15.5Na-citrate - - - 20Na-gluconate - 40 - -Na-pyruvate 100 - - -Citric acid - - - 2Inosine 100 - - -Adenine 5 1.4 1.25 2Guanosine - 1.4 - -Glucose - 47 45 61Mannitol - 55 30 -pH 6.7-7.4 8.2 6.2 5.8Osmolality (mOsm/kg) 510 275 376 291


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guidelines and practices. This study was approved by the institutional medical ethical committees, in accordance with the standards laid down in the 1964 Declaration of Helsinki.

Whole blood (WB), 500 mL ± 2%, was collected in quadruple bags, bottom-and-top collection systems containing 70 mL of CPD (Fresenius Kabi, Emmer Compascuum, the Netherlands). The day of blood collection was designated as day 0 of the study. The WB units were placed on butane1,4-diol cooling plates (Compocool, Fresenius Kabi) to allow their temperatures to adjust to 20 to 24°C.25 After overnight hold, the WB units were processed according to the routine buffy coat procedure as previously described.26 Briefly, after a hard spin (Sorvall RC12BP), the WB units were separated in components, using an automated blood component separator (Compomat G5, Fresenius HemoCare). After separation, 110 mL of additive solution, SAGM or PAGGGM, was added to the RBC via the filter. After careful mixing the units were WBC reduced using the inline leukoreduction filter (BioR, Fresenius Kabi). The RBC units were subsequently stored at 2-6°C for seven days prior to rejuvenation (if applicable), glycerolization and freezing.

RejuvenationThe rejuvenation procedure with Rejuvesol (Citra Labs, Braintree, MA) was carried out according to the instructions of the manufacturer. Units meant to be rejuvenated were incubated in a water bath, maintained at 37°C, for 60 min with 50 mL of Rejuvesol, containing 0.550 g of sodium pyruvate, 1.340 g of inosine, 0.034 g of adenine, 0.730 g of dibasic sodium phosphate and 0.311 g of monobasic sodium phosphate, in water for injection, with an osmolality of 510 mOsm/kg H2O and pH 6.7-7.4. (Table 1)

GlycerolizationRBC units were centrifuged at 3200 x g for 5 minutes. The additive solution was removed to produce RBC units with a hematocrit (Hct) of 75 ± 5%. The RBC units were sterilely connected (TSCD Terumo-BCT, Lakewood, CO) to a disposable glycerolization set (LN225, Haemonetics) and glycerolization was completed using the ACP215. A volume of 6.2 mol/L (57%) glycerol solution (S.A.L.F. S.p.A., Cenate Sotto, Bergamo, Italy) was added to the red cells to achieve a final glycerol concentration of 40% (w/v). The glycerolized cells were transferred to a 1000 mL PVC bag (VSE 6002Z, MacoPharma, Mouvaux,

Figure 1. Study design, time frame and flow diagram.

Series I. At day 8 after collection, 4 units of RBC in SAGM were pooled (n=4), mixed and split. From each pool

two units were incubated with Rejuvesol before freezing. All units were glycerolized within one hour after

rejuvenation, frozen, stored for at least 14 days at -80°C and deglycerolized. From each pair, one unit was

resuspended in SAGM, the other in AS-3. All thawed units were subsequently stored at 2-6 ⁰ C for at least 35

days.

Series II. At day 8 after collection, 2 units of RBC in PAGGGM were pooled (n=4), mixed and split. All units were

then glycerolized, frozen, stored for at least 14 days at -80°C and deglycerolized. From each pair, one unit was

resuspended in SAGM, the other in AS-3. All thawed units were subsequently stored at 2-6 ⁰ C for at least 35

days.

Series I Series II

RBC in AS-3

RBC in SAGM

RBC in AS-3

RBC in SAGM

RBC in AS-3

RBC in SAGM

Glycerolization, frozen storage, deglycerolization

Rejuvenation

RBC in SAGM

RBC in SAGM

RBC in SAGM

RBC in SAGM

Day 8: Pool in SAGM

RBC in SAGM

RBC in SAGM

RBC in SAGM

RBC in SAGM

RBC in PAGGGM

RBC in PAGGGM

RBC in PAGGGM

M

RBC in PAGGGM

M

Day 8: Pool in PAGGGM

Glycerolization, frozen storage, deglycerolization

Figure 1. Study design, time frame and flow diagram. Series I.

At day 8 after collection, 4 units of RBC in SAGM were pooled (n=4), mixed and split. From each pool two units were incubated with Rejuvesol before freezing. All units were glycerolized within one hour after rejuvenation, frozen, stored for at least 14 days at -80°C and deglycerolized. From each pair, one unit was resuspended in SAGM, the other in AS-3. All thawed units were subsequently stored at 2-6⁰ C for at least 35 days.

Series II. At day 8 after collection, 2 units of RBC in PAGGGM were pooled (n=4), mixed and split. All units were then glycerolized, frozen, stored for at least 14 days at -80°C and deglycerolized. From each pair, one unit was resuspended in SAGM, the other in AS-3. All thawed units were subsequently stored at 2-6⁰ C for at least 35 days.


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France) The RBC units were sealed in a plastic overwrap and frozen as described before.27 All frozen RBC units were stored at -80°C for at least 14 days.

Thawing, deglycerolization and storageThe frozen RBCs were thawed in a water bath maintained at 40°C, to achieve a surface temperature of 25-30°C as measured with a non-contact thermometer (Fluke 62 mini, Eindhoven, the Netherlands). The thawed RBC units were sterilely connected to a deglycerolization set (LN235, Haemonetics). The thawed RBC units were washed in the ACP215 using 50 mL of 12% NaCl (Bio-Deglyc, Bioluz, Saint-Jean-de-Luz, France) followed by several rounds of washing with in total 1.6 L of Phosphate Buffered Saline (PBS, pH 7.4).22 Upon completion of the deglycerolization process, the cells were washed with, and finally resuspended in either SAGM (Haemonetics) or AS-3 (Haemonetics). The deglycerolized RBC units were planned to be stored at 2-6°C for at least 35 days.

The efficacy of the deglycerolization procedure was assessed by the measurement of supernatant osmolality.28 Supernatant osmolality of the deglycerolized units was measured using an Osmomat 030-D Cryoscopic osmometer (Gonotec, Berlin, Germany), immediately after the deglycerolization procedure.

Hematological parameters and recovery The volume of the blood components was calculated from the net weight divided by the specific gravity: 1.027 g/mL for plasma, 1.100 for 40% glycerol and RBCs, and 1.006 for SAGM. Using the hematocrit of the solution, the specific gravity of RBC in additive solution was calculated from these values. Freeze-thaw-wash recovery was calculated by comparing the total Hb after deglycerolization to the total Hb prior to glycerolization. Total Hb was calculated as volume (L) x Hb concentration (g/L). Hb concentration and red blood cell count were determined on a hematology analyzer (Advia 2120, Siemens Medical Solutions Diagnostics, Breda, the Netherlands). Hct was measured with the spun capillary method. Hemolysis was determined as described previously.23 Briefly, cell-free supernatants were obtained by centrifugation of the RBC suspensions at 12000 x g for 5 min followed by an additional centrifugation of the supernatant at 12000 x g for 30 sec. The used centrifugation protocol was checked upon loss of microparticles (formed during storage) from the supernatant. It was found that microvesicles were not significantly removed.

Free Hb was determined by absorbance measurement of supernatant at 415 or 514 nm by a spectrophotometer (Eon plate reader, Bio Tek, Bad Friedrichshall, Germany), with correction for plasma absorption if necessary. Hemolysis was expressed as a percentage of total Hb present in the RBC after correction for hematocrit.

Analysis of metabolic and cellular variables Extracellular potassium, glucose, lactate and pH were measured with a blood gas analyzer (Rapidlab 1265, Siemens Medical Solutions Diagnostics).

2,3-DPG and adenine nucleotides (ATP, ADP, AMP) levels were determined in neutralized perchloric acid extracts. 2,3-DPG was analyzed with an enzymatic assay from Roche (Mannheim, Germany). Adenine nucleotides were assayed, using a HPLC method.29 The measured concentrations (μmol/L) were divided by the Hb concentration (g/L). Values for 2,3-DPG and adenine nucleotides are thus expressed in μmol/g Hb. Total adenylate content was calculated as the sum of ATP, ADP and AMP levels.

PS-exposureTo quantify the amount of erythrocytes exposing phosphatidylserine (PS) on their cell surface, cells were stained with FITC-labelled Annexin V, essentially according to Verhoeven et al.6 Briefly, samples of red cells were washed twice with an incubation medium consisting of 134 mM NaCl, 10 mM glucose, 10 mM Tris-HCl, and 40 mM sucrose, 20 mM Hepes and 0.5% human serum albumin (HSA) (pH 7.4). After washing, the cells were resuspended at 0.3% Hct in the same medium supplemented with 2.5 mM CaCl2. Labeling with Annexin V was performed by adding Annexin V-FITC (VPS-Diagnostics, Hoeven, the Netherlands) to a final concentration of 1 μg/ml to 250 μL cell suspension. After incubation at room temperature (RT) in the dark for 30 min, cells were washed once and analyzed on a flow cytometer (LSRII Becton Dickinson, Breda, the Netherlands). The percentage of Annexin V positive cells was determined by comparison with a negative control that was incubated with EGTA.

DeformabilityThe deformability of a red cell is an indication of its capability to deform during its passage through the narrowest capillaries of the microcirculation and thus its capacity to survive after transfusion. Expressed as the deformation index (DI),


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the deformability was defined as the ratio of the major axis length to the minor axis width. Thus, the DI of a disc shaped red cell by definition is equal to 1.

The DI was determined with an Automated Rheoscope and Cell Analyzer (ARCA, Mechatronics Instruments, Hoorn, the Netherlands).30 In short, a thin layer of RBC is sheared between two horizontal plates, one of which is able to rotate with variable speed at an adjustable distance from the other. These variables lead to a variable, deforming force exerted on the red cells. An increasing force results in elongation (deformation) of the RBC in the resulting current. This process is measured by a laser beam diffraction pattern, captured on a video camera and analyzed by a computer on individual cell basis. Results are expressed as percentage of cells with DI <1.5 (non-deformable cells) and DI >2.5 (deformable cells).

Statistical analysisResults are shown as mean ± SD. Results obtained after pf-R (paired) or pf-PAGGGM (unpaired) were compared with the non-rejuvenated cells in SAGM (control). Results obtained during postthaw storage after pf-R (paired) and pf-PAGGGM (unpaired) were compared with the non-rejuvenated cells (control) in the same postthaw additive solution. Results were analyzed with repeated-measures analysis of variance (ANOVA), with a Dunnett’s post test to compare the values with (non-rejuvenated) control cells (Instat, Version 3.06, GraphPad, San Diego, CA). Differences were considered significant when p values were less than 0.05.

Results

Effects of Rejuvesol and PAGGGMCompared to control, pf-R increased the ATP levels of RBC stored for 7 days in SAGM, from 5.5 ± 0.1 μmol/g Hb to 7.5 ± 0.3 μmol/g Hb (a mean increase of 36%)(Table 2), partly at the expense of ADP (decrease from 0.84 ± 0.12 to 0.30 ± 0.02 µmol/g Hb). Total adenylate levels rose from 6.5 ± 0.1 μmol/g Hb to a mean of 8.2 ± 0.2 μmol/g Hb (a mean increase of 27%) in the pf-R (Table 2). In the pf-PAGGGM units, we measured a mean ATP and a mean total adenylate content of 6.2 ± 0.4 μmol/g Hb and 6.8 ± 0.3 μmol/g Hb respectively, indicating a small positive effect, compared to the (non-rejuvenated) control units in SAGM (Table 2).

Levels of 2,3-DPG in the pf-R climbed from 4.6 ± 2.1 μmol/g Hb to 22.7 ± 1.7 μmol/g Hb (a mean increase of almost 400%). The pf-PAGGGM units showed a level of 14.7 ± 1.6 μmol/g Hb (Table 2).

Freezing/Thawing/Deglycerolization characteristicsHemoglobin content of the RBC units before freezing was comparable, indicating efficient pooling and splitting. Volumes of the rejuvenated RBCs were higher because of the addition of Rejuvesol (Table 2).

After thawing, the volume of all units was around 300 mL with a hemoglobin content of around 40 g. The freeze-thaw-wash recovery was around 80%, with no difference between the different groups (Table 2).

Table 2. Characteristics of starting products (SAGM and PAGGGM) and deglycerolized products (SAGM and AS-3)

SAGM (Non-rejuvenated)

SAGM (Rejuvenated)

PAGGGM

Starting productVolume (mL) 274 ± 11.3 310 ± 9.6* 277 ± 12.3Total Hb (g) 50.2 ± 2.8 49.8 ± 2.4 50.7 ± 2.2ATP 5.5 ± 0.1 7.5 ± 0.3* 6.2 ± 0.4*Total Adenylate 6.5 ± 0.1 8.2 ± 0.2* 6.8 ± 0.32,3 DPG 4.6 ± 2.1 22.7 ± 1.7* 14.7 ± 1.6*

Deglycerolized productVolume (mL) 298 ± 2.4 300 ± 1.9 301 ± 1.6Total Hb (g) 40.0 ± 2.7 40.2 ± 2.7 40.8 ± 1.7Hct in SAGM (%) † 43 ± 4.2 42 ± 4.0 41 ± 2.6Hct in AS-3 (%) † 50 ± 3.6* 46 ± 5.0 48 ± 2.1

Freeze-Thaw-Wash recovery (%) 80 ± 4 81 ± 4 80 ± 3

SAGM AS-3Supernatant osmolality (mOsm/kg)# 345 ± 14 302 ± 3*pH (at 37°C)# 6.72 ± 0.08 6.49 ± 0.03*

Data are reported as mean ± SD (n=8), †: n=4, #: n=12*: p < 0.05 as compared to SAGM


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Hematocrit of the thawed units depended on the additive solution used, being higher with AS-3 compared to SAGM. Also pH depended on the additive solution, being lower for AS-3 than SAGM (Table 1 and 2).

The osmolality of the supernatant of all 24 deglycerolized units was well below 420 mOsm/kg and comparable for the three groups (Table 2), indicating adequate removal of glycerol (remaining glycerol level less than 1%).1,19 The osmolality of the supernatant of the SAGM units was considerably higher than of the AS-3 units (Table 2), due to the difference in osmolality of these two additive solutions (Table 1).

Postthaw storage characteristicsHemolysisUnits stored in SAGM showed clearly higher hemolysis levels than the AS-3 units (Fig. 2). Already at day 14 of postthaw storage in SAGM, hemolysis reached levels above the 0.8% (European) threshold. In AS-3, postthaw hemolysis remained below 0.8% for at least 35 days. Because of the favorable results obtained with AS-3, the measurements in AS-3 units were continued till day 42 (Fig. 2-4). Prefreeze rejuvenation or incubation in PAGGGM had no influence on postthaw hemolysis. ATP and Total AdenylateImmediately after thawing, ATP levels were increased in both the pf-R and the pf-PAGGGM (Fig. 3). During storage, ATP levels gradually decreased, with the decrease being faster in the pf-R. During postthaw storage in SAGM, ATP levels of pf-R remained significantly higher than of the non-rejuvenated cells during the first two weeks of storage only, whereas for pf-PAGGGM, ATP levels were higher during 35 days of storage.

In AS-3, ATP-levels of both pf-R and pf-PAGGGM were significantly higher than of the (non-rejuvenated) control cells for 35 days.

Total adenylate levels were increased due to the treatment with Rejuvesol before freezing and remained significantly higher than in the controls during the whole postthaw storage time, both in SAGM and AS-3 (Fig. 4). The same was true for cells prefreeze stored in PAGGGM. At the end of the storage period, (day 35 in SAGM, day 42 in AS-3), total adenylate levels in pf-R and pf-PAGGGM were comparable or even slightly higher as compared to (non-rejuvenated) control cells immediately after thawing (Fig. 4).

A

B

Figure 2. Hemolysis in deglycerolized red cells during postthaw storage in SAGM (A) and AS-3 (B), with prefreeze rejuvenation (pf-R, C), PAGGGM (pf-PAGGGM, MCM) or control (FCF). Results shown represent mean ± SD (n=4).


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Figure 3. ATP content in deglycerolized red cells during postthaw storage in SAGM (A) and AS-3 (B), with prefreeze rejuvenation (pf-R, C), PAGGGM (pf-PAGGGM, MCM) or control (FCF). Results shown represent mean ± SD (n=4). *: p < 0.05 as compared to control.

A

B

A

B

Figure 4. Total adenylate content in deglycerolized red cells during postthaw storage in SAGM (A) and AS-3 (B), with prefreeze rejuvenation (pf-R, C), PAGGGM (pf-PAGGGM, MCM) or control (FCF). Results shown represent mean ± SD (n=4). *: p < 0.05 as compared to control.


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2,3-DPGDuring the first 2 weeks of postthaw storage, 2,3-DPG levels of pf-R and pf-PAGGGM were significantly higher as compared to control cells, with only small differences between storage in SAGM and AS-3 (Fig. 5). At day 21 of storage, 2,3-DPG of all units was around the limit of detection. Despite the higher initial levels in pf-R, at day 14 of storage, 2,3-DPG levels were comparable for pf-PAGGGM and pf-R.

PS- exposure and deformabilityAt day 35 of storage, the percentage of Annexin V positive cells amounted to 4.7 ± 1.6% in SAGM and was significantly higher (7.9 ± 2.5%) when stored in AS-3 (Table 3). PS-exposure was not influenced by pf-R or pf-PAGGGM, regardless of the postthaw additive solution. Deformability was measured at day 35 of postthaw storage. pf-PAGGGM showed a higher percentage of non-deformable cells (DI <1.5) and a lower percentage of deformable cells (DI >2.5) as compared to control or pf-R. This effect was significantly more pronounced during storage in AS-3 as compared to storage in SAGM. For the control and pf-R, there were no significant differences between storage in AS-3 and SAGM (Table 3).

A

B

Figure 5. 2,3-DPG content in deglycerolized red cells during postthaw storage in SAGM (A) and AS-3 (B), with prefreeze rejuvenation (pf-R, C), PAGGGM (pf-PAGGGM, MCM) or control (FCF). Results shown represent mean ± SD (n=4). *: p < 0.05 as compared to control.

Table 3. Annexin V positive cells and Deformability Index (DI) after 35 days of postthaw storage

Prefreeze treatment Postthaw additive solution

% Annexin V pos cells

DI <1.5 DI >2.5

None SAGM 4.6 ± 0.4 1.9 ± 0.4 76 ± 5.7pf-R SAGM 5.5 ± 2.0 2.2 ± 1.2 67 ± 4.5pf-PAGGGM SAGM 4.1 ± 1.9 3.1 ± 0.9 64 ± 5.3*None AS-3 8.2 ± 1.5 3.4 ± 1.5 63 ± 11pf-R AS-3 7.7 ± 1.5 3.2 ± 1.5 63 ± 9.8pf-PAGGGM AS-3 7.9 ± 4.4 6.3 ± 1.1* 56 ± 1.9*

Results shown represent mean ± SD (n=4). *: p < 0.05 as compared to control (no prefreeze treatment) in same additive solution.


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Discussion

Currently, US and European guidelines1,3 both allow for thawed, deglycerolized, red cells to be stored for longer than 24 hrs., if prepared in a closed system. The AABB1 and FDA2 even specifically mention a postthaw storage expiration time of 14 days, enabled by the use of AS-3 as additive solution.31 Regardless of the expiration time, the maximum allowable hemolysis at the time of infusion is 1.0% in the US1 and 0.8% in Europe.3 Furthermore, both the US and Europe require a 24 hr posttransfusion survival of at least 75%.3,32 Obviously, any further extension of this postthaw shelf life, while still meeting all the previous requirements, would simplify frozen red cell inventory management by limiting unnecessary wastage and shortages, from which both civilian and military transfusion practice could benefit.

Particularly hemolysis and energy status are of importance when looking at possibilities to extend the currently approved shelf life. Previous adjustments in the glycerolization and washing procedure20,22 helped to achieve a postthaw shelf life of 28 days in AS-3. Whereas hemolysis stayed below 0.8% for even 35 days, the energy status, as measured by the total adenylate content, proved to be the limiting factor.

ATP content is indirectly related to posttransfusion survival, because normal ATP levels are necessary to prevent membrane loss by microvesiculation and also to maintain active, outward transport of phospholipids like phosphatidylserine (PS), thus preventing premature clearance from the recipient’s circulation by macrophages.33

Rejuvesol and the recently developed alkaline, chlorine-free additive solution PAGGGM23,24 are known to be able to increase ATP (and also 2,3-DPG) levels. The inosine and adenine in Rejuvesol are the key factors in raising the levels of 2,3-DPG and ATP respectively via the Embden-Meyerhof (glycolytic) pathway.34 Most likely, PAGGGM has a positive effect on the activity of phosphofructokinase (PFK) in the glycolytic pathway, thus raising the levels of both ATP and 2,3-DPG.24

Prefreeze rejuvenation of red cells stored for 7 days at 2-6°C in SAGM, and, to a lesser extent, the replacement of SAGM by PAGGGM, resulted in significantly higher levels of 2,3-DPG, ATP and total adenylate (Table 2). These increased levels were maintained during the frozen storage and the deglycerolization procedure.

Although ATP levels were better maintained during postthaw storage in SAGM compared to storage in AS-3, SAGM failed to keep hemolysis below 0.8% after 7 days in all units (Fig. 2A). If resuspended in AS-3, however, for both (non-rejuvenated) control, pf-R and pf-PAGGGM, hemolysis stayed below the threshold of 0.8% up to 35 days, with no differences between the different groups (Fig. 2B). This lower hemolysis in AS-3, as compared to SAGM, appears to be completely attributable to its citrate content.22,35 This not only confirms our earlier results with regard to hemolysis,20,22 but also the superiority of AS-3 as a long-term storage medium for deglycerolized red cells.

During postthaw storage, the ATP levels gradually declined, accompanied by an increase in the levels of ADP and AMP. In the glycolytic pathway, both AMP and ADP can be converted back to ATP, so both these substances still contribute to the cell’s energy status, expressed as the total adenylate content. Only when AMP is converted to IMP, which is irreversible for erythrocytes, the potential to recover ATP is lost, but no increase in IMP was found (data not shown). US and European guidelines dictate an “in vivo” 24 hr posttransfusion survival of at least 75%.1,3,32 As a surrogate marker for “in vivo” survival, total adenylate content could be used. Högman et al.36 not only found a good correlation between total adenylate content and posttransfusion recovery, but also that the minimum total adenylate content needed to meet this requirement was 82% of the original levels. Whereas (non-rejuvenated) control units failed to meet this requirement at day 35, all pf-R and pf-PAGGGM showed total adenylate levels above this level, independent of the additive solutions used for postthaw storage (results not shown). So, in contrast to our earlier study in which total adenylate content limited the postthaw storage,20 hemolysis was the limiting factor in pf-R and pf-PAGGGM during postthaw storage in AS-3.

Not surprisingly, because of the 7 days storage, the (non-rejuvenated) control units showed 2,3-DPG levels already close to zero before freezing (Fig. 5). During the first 14 days of postthaw storage in both SAGM and AS-3, pf-R and pf-PAGGGM showed significantly higher levels of 2,3-DPG (Fig. 5), suggesting better oxygen delivery capacity, from which certain patients may benefit.11,18,23 Levels of 2,3-DPG at day 7 in both SAGM and AS-3 were still comparable to the ones of fresh red cells at the beginning of routine (refrigerated) blood bank storage at 2-6°C.5

We postulate that the acidic postthaw environment, caused by SAGM and even more by AS-3, quickly neutralized the favorable effect of Rejuvesol


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and PAGGGM on 2,3-DPG levels during storage. Already from day 0 on, pH in all units was well below the turning point of 7.2 (Table 2), at which biphosphoglycerate phosphatase is activated,37 causing a rapid loss of 2,3-DPG.38 At the same time, due to the lower pH of SAGM and AS-3, the prevailing effect of bisphosphoglycerate mutase on the Rapoport-Luebering shunt sustains the production of ATP further down the glycolytic pathway. This would explain the longer lasting effect of Rejuvesol and PAGGGM on ATP levels in the deglycerolized units.

Storage of leukoreduced red cells in routine refrigerated blood banking shows very low levels of PS-exposure, as measured by the number of Annexin V positive cells, even after 35 days.6 Cryopreservation by itself does not cause PS-exposure.39 The higher percentages of PS-exposure we found at day 35 in all units, may partly be attributable to the longer prefreeze storage time,39 but especially for the AS-3 units the lower pH of around 6.49 (Table 2) could be responsible.40

Deformability is strongly related to normal red cell function and red cell viability,41 but in itself is not adversely affected by cryopreservation and the subsequent thaw and deglycerolization procedure.42 Rejuvesol-treated red cells, stored under normal, refrigerated, blood bank conditions at 2-6°C, are reported to have an improved elasticity.14 In our study, prefreeze treatment with Rejuvesol did not lead to lower percentages of red cells with impaired deformability, as determined by a Deformability Index (DI) < 1.5. On the other hand, prefreeze storage in PAGGGM did result in a slightly higher percentage of non-deformable cells during postthaw storage.

The levels of ATP in AS-3 resuspended red cells at day 35, although on average below 2.5 μmol/g Hb, did not show the correlation with the deformability results, as reported by Karger et al.43 Although speculative, based on the total adenylate content and its predictive value considering 24 hr posttransfusion survival, deformability defects may very well be restored upon transfusion, thus limiting possible negative rheological effects at the microcirculatory level.

The rather long prefreeze storage time of 7 days, chosen because of logistical reasons, as well as the low pH of AS-3, have most probably negatively influenced the outcome of this study, particularly with regard to 2,3-DPG, since this substance is strongly affected by length of prefreeze storage at 4°C.39,44 Furthermore, maintaining a physiological intracellular pH of around 7.4 by at least raising the pH of the final resuspension medium in particular, with citrate as an important

constituent, would also potentially help to improve future results. Even better results may probably be obtained by performing the whole procedure from collection to postthaw storage at alkaline levels, as proposed in earlier studies on processing and storage of liquid red cells.23,24,38,45,46 In this respect, PAGGGM, with its pH of 8.0, could be the additive solution of choice. Currently, PAGGGM is not CE-marked, but only designed and used for research purposes. Further research on the use of PAGGGM as additive solution is planned, including “in vivo” evaluation of erythrocytes stored in PAGGGM. However, since for maintaining postthaw stability the presence of citrate seems to be necessary, PAGGGM cannot be used for long-term storage of thawed cells. A PAGGGM-based solution, with (part of) the mannitol replaced by citrate, might give both the desired postthaw stability and high levels of intracellular organic phosphate compounds.

In conclusion, our current findings indicate the possibility of extending the postthaw shelf life of red cells, rejuvenated with Rejuvesol or prefreeze stored in PAGGGM, to at least 35 days. The important practical implication of this study is that it facilitates operating a frozen blood bank, by minimizing the effects of a time consuming deglycerolization procedure. Thus, the ability to maintain a liquid, previously frozen, inventory of red cells creates the possibility to better meet unexpected operational demands, as is the case under battle field conditions e.g. Operational and logistical advantages may thus very well outweigh the higher cost of maintaining and operating a frozen blood bank, including the additional cost of using Rejuvesol or PAGGGM.


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References

1. AABB Standards for Blood Banks and Transfusion Services. 30th edition, 2016.2. FDA Circular of Information for the Use of Human Blood and Blood Components,

revised November 2013, 2016.3. Council of Europe. Guide to the Preparation, Use and Quality Assurance of Blood

Components. 18th edition, 2015. Strasbourg 4. De Korte D, Verhoeven AJ. Quality determinants of erythrocyte destined for

transfusion. Cell Mol.Biol.(Noisy.-le-grand) 2004;50(2):187-95.5. Hess JR. Measures of stored red blood cell quality. Vox Sang. 2014;107(1):1-9.6. Verhoeven AJ, Hilarius PM, Dekkers DWC, Lagerberg JWM, de Korte D. Prolonged

storage of red blood cells affects aminophospholipid translocase activity. Vox Sang. 2006;91(3):244-51.

7. Longster GH, Buckley T, Sikorski J, Derrick Tovey LA. Scanning electron microscope studies of red cell morphology. Changes occurring in red cell shape during storage and post transfusion. Vox Sang. 1972;22(2):161-70.

8. Greenwalt TJ, Bryan DJ, Dumaswala UJ. Erythrocyte membrane vesiculation and changes in membrane composition during storage in citrate-phosphate-dextrose-adenine-1. Vox Sang. 1984;47(4):261-70.

9. van de Watering LMG, Brand A. Effects of storage of red cells. Transfus.Med Hemother. 2008;35(5):359-67.

10. Offner PJ. Age of blood: does it make a difference? Crit Care 2004;8 Suppl 2:S24-S26.

11. Valeri CR. Use of rejuvenation solutions in blood preservation. Crit Rev.Clin.Lab Sci. 1982;17(4):299-374.

12. Usry RT, Moore GL, Manalo FW. Morphology of stored, rejuvenated human erythrocytes. Vox Sang. 1975;28(3):176-83.

13. Raat NJH, Hilarius PM, Johannes T, de Korte D, Ince C, Verhoeven AJ. Rejuvenation of stored human red blood cells reverses the renal microvascular oxygenation deficit in an isovolemic transfusion model in rats. Transfusion 2009;49(3):427-4.

14. Gelderman MP, Vostal JG. Rejuvenation improves roller pump-induced physical stress resistance of fresh and stored red blood cells. Transfusion 2011;51(5):1096-104.

15. Koshkaryev A, Zelig O, Manny N, Yedgar S, Barshtein G. Rejuvenation treatment of stored red blood cells reverses storage-induced adhesion to vascular endothelial cells. Transfusion 2009;49(10):2136-43.

16. Valeri CR, Zaroulis CG. Rejuvenation and freezing of outdated stored human red cells. N.Engl.J.Med. 1972;287(26):1307-13.

17. Moore GL, Failla ML, Blake BH, Gray JL, Manalo FW. Storage rejuvenation, and restorage of human cells for periods up to 56 days. Transfusion 1974;14(3):249-52.

18. Dennis RC, Hechtman HB, Berger RL, Vito L, Weisel RD, Valeri CR. Transfusion of 2,3 DPG-enriched red blood cells to improve cardiac function. Ann.Thorac.Surg. 1978;26(1):17-6.

19. Clinical and Practical Aspects of the Use of Frozen Blood. AABB; 1977.pp 23-36.20. Lelkens CCM, de Korte D, Lagerberg JWM. Prolonged post-thaw shelf life of red cells

frozen without prefreeze removal of excess glycerol. Vox Sang. 2015;108(3):219-25.



23. De Korte D, Kleine M, Korsten HGH, Verhoeven AJ. Prolonged maintenance of 2,3-diphosphoglycerate acid and adenosine triphosphate in red blood cells during storage. Transfusion 2008;48(6):1081-9.

24. Burger P, Korsten H, de Korte D, Rombout E, Van Bruggen R, Verhoeven AJ. An improved red blood cell additive solution maintains 2,3-diphosphoglycerate and adenosine triphosphate levels by an enhancing effect on phosphofructokinase activity during cold storage. Transfusion 2010;50(11):2386-92.


26. Van der Meer P, Pietersz R, Hinloopen B, Dekker W, Reesink H. Automated separation of whole blood in top and bottom bags into components using the Compomat G4. Vox Sang. 1999;76(2):90-9.


28. Valeri CR, Pivacek LE, Cassidy GP, Ragno G. The survival, function, and hemolysis of human RBCs stored at 4 degrees C in additive solution (AS-1, AS-3, or AS-5) for 42 days and then biochemically modified, frozen, thawed, washed, and stored at 4 degrees C in sodium chloride and glucose solution for 24 hours. Transfusion 2000;40(11):1341-5.

29. De Korte D, Haverkort WA, van Gennip AH, Roos D. Nucleotide profiles of normal human blood cells determined by high-performance liquid chromatography. Anal.Biochem. 1985;147(1):197-209.

30. Dobbe JGG, Streekstra GJ, Hardeman MR, Ince C, Grimbergen CA. Measurement of the distribution of red blood cell deformability using an automated rheoscope. Cytometry 2002;50(6):313-25.


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32. Dumont LJ, AuBuchon JP. Evaluation of proposed FDA criteria for the evaluation of radiolabeled red cell recovery trials. Transfusion 2008;48(6):1053-60.

33. Kamp D, Sieberg T, Haest CW. Inhibition and stimulation of phospholipid scrambling activity. Consequences for lipid asymmetry, echinocytosis, and microvesiculation of erythrocytes. Biochemistry 2001;40(31):9438-46.

34. Meyer EK, Dumont DF, Baker S, Dumont LJ. Rejuvenation capacity of red blood cells in additive solutions over long-term storage. Transfusion 2011;51(7):1574-9.

35. Besselink GAJ, Ebbing IG, Hilarius PM, de Korte D, Verhoeven AJ, Lagerberg JWM. Composition of the additive solution affects red blood cell integrity after photodynamic treatment. Vox Sang. 2003;85(3):183-9.


37. Hess JR, Greenwalt TG. Storage of red blood cells: new approaches. Transfus.Med.Rev. 2002;16(4):283-95.

38. Hogman CF, Knutson F, Loof H, Payrat JM. Improved maintenance of 2,3 DPG and ATP in RBCs stored in a modified additive solution. Transfusion 2002;42(7):824-9.

39. Holovati JL, Wong KA, Webster JM, Acker JP. The effects of cryopreservation on red blood cell microvesiculation, phosphatidylserine externalization, and CD47 expression. Transfusion 2008;48(8):1658-68.

40. Libera J, Pomorski T, Muller P, Herrmann A. Influence of pH on phospholipid redistribution in human erythrocyte membrane. Blood 1997;90(4):1684-93.

41. Hogman CF, Meryman HT. Red blood cells intended for transfusion: quality criteria revisited. Transfusion 2006;46(1):137-42.

42. Henkelman S, Lagerberg JWM, Graaff R, Rakhorst G, Van Oeveren W. The effects of cryopreservation on red blood cell rheologic properties. Transfusion 2010;50(11):2393-401.

43. Karger R, Lukow C, Kretschmer V. Deformability of Red Blood Cells and Correlation with ATP Content during Storage as Leukocyte-Depleted Whole Blood. Transfus.Med Hemother. 2012;39(4):277-82.

44. Valeri CR, Ragno G, Van Houten P, Rose L, Rose M, Egozy Y, Popovsky MA. Automation of the glycerolization of red blood cells with the high-separation bowl in the Haemonetics ACP 215 instrument. Transfusion 2005;45(10):1621-7.

45. Guppy M, Attwood PV, Hansen IA, Sabaratnam R, Frisina J, Whisson ME. pH, temperature and lactate production in human red blood cells: implications for blood storage and glycolytic control. Vox Sang. 1992;62(2):70-5.

46. Hess JR, Hill HR, Oliver CK, Lippert LE, Greenwalt TJ. Alkaline CPD and the preservation of RBC 2,3-DPG. Transfusion 2002;42(6):747-52.

John R. Hess1, Charles C.M. Lelkens2, John B. Holcomb3 and Thomas M. Scalea4

1 Department of Laboratory Medicine, University of Washington, Seattle, USA 2 Royal Netherlands Navy, Medical Corps, retired, Kortgene, the Netherlands

3 Division of Acute and Critical Care Surgery, University of Texas at Houston, Houston, USA 4 The Program in Trauma, University of Maryland School of Medicine, Baltimore, USA

Transfus Apher Sci. 2013 Dec; 49(3):380-6.

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Advances in military, field, and austere transfusion medicine in the last decade

7

Chapter 7 Advances in military, field, and austere transfusion medicine in the last decade

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Abstract

Two decades of war in Southwest Asia has demonstrated the essential role of primary resuscitation with blood products in the care of critically injured soldiers. This idea has been widely adopted and is being critically tested in civilian trauma centers. The need for red cells, plasma and platelets to be immediately available in remote locations creates a logistic burden that will best be eased by innovative new blood products such as longer-stored liquid RBCs, freeze-dried plasma, small-volume frozen platelets, and coagulation factor concentrates such as fibrinogen concentrates and prothrombin complex concentrates. Such products have long shelf lives, low logistic burdens of weight, fragility, or needs for processing prior to use. Developing and fielding a full family of such products will improve field medical care and make products available in the evacuation chain. It also will allow treatment in other austere environments such as the hundreds of small hospitals in the US which serve as Levels 3 and 4 trauma centers but do not currently have thawed plasma or platelets available. Such small trauma centers currently care for half of all the trauma patients in the country. Proving the new generation of blood products work, will help assure their widest availability in emergencies.

Introduction

Military transfusion medicine is the specialty of developing, deploying, and using blood products for medical care in austere combat environments and in the medical evacuation chain. Field transfusion medicine is the civilian equivalent used in disaster planning and relief. Field practices are relevant in other resource-limited settings such as small or midsized hospitals that do not have all blood products available as, in the US, 50% of trauma patients are cared for outside of Levels 1 and 2 trauma centers.

Interplay is frequent between military and civilian field care because the military has a mission to support disaster relief. The military has assets such as aircraft, prepackaged medical equipment, trained deployable surgical teams, deployable hospitals, and theater blood transshipment facilities and treaties to carry out such missions internationally. As a result, military casualty care research programs have provided much of the epidemiologic data that has supported field blood-use doctrine and have paid for most of the development of modern blood storage systems. Retired military personnel are often used as experts in disaster planning. Shared equipment, training, experience, doctrine, and literature have all contributed to an evolving sense of best field medical practices over the last decade.1

The military often plans from the experience of prior wars, and so the first US invasion of Iraq in 1990 is instructive.2 Multiple field hospitals were deployed along with 82,000 units of packed red blood cells (RBC). In hindsight, the 250 US casualties and 250 units of RBC used in their care in the whole war could have been handled with a few smaller combat support hospitals and a dozen cardboard and styrofoam boxes of blood products. However, the critical casualty of the war sustained a transpelvic fragment wound and required 52 units of RBCs and ultimately fresh whole blood to treat his iatrogenic dilutional coagulopathy. The lessons that blood use at a per casualty rate is generally low but that the patients receiving massive transfusions require more than just red cells were confirmed in the actions in Somalia, Bosnia, and Kosovo. As US military budgets got smaller at the end of the decade, many large deployable military field hospitals were deactivated and small forward surgical teams were redeveloped to provide immediate far forward care. Such teams could carry 20 units of RBCs on ice and blood bags for the collection of fresh whole blood from soldiers if plasma or platelets were needed, but the forward surgical teams did not have the assets to store frozen plasma or platelets.


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Also in the 1990s, changes in the education of acute care (trauma) surgeons followed evolving theory and improving techniques. Early in this decade, ‘‘damage control surgery’’ was defined as an approach to stabilize patients whose injuries were too numerous or severe to be repaired in a single survivable procedure.3 Damage control involved quick hemorrhage control by vascular shunting and organ and soft tissue packing and management of body cavity contamination by tying off gut and diverting bile and urine. These actions saved lives, but created patients whom the military did not know if they could safely transport. Surgeons had learned to save a group of patients that previously died, but were now too sick for prolonged care in the austere environment. Transport of such patients out of the austere environment must occur early, in a window of relative stability. US Air Force medical evacuation personnel were first exposed to these patients in Somalia in 1993 and began developing critical care air transport teams to manage them.4 Prolonged critical care of these most seriously injured is a profound logistic burden in even the best Level 1 trauma centers.

In the second half of the decade, efforts to prevent acute respiratory failure and compartment syndromes by reducing non-blood fluid administration achieved notable successes in clinical trials in the academic settings where the military trained its trauma surgeons.5,6

Nevertheless, academic specialty groups like the Committee on Trauma of the American College of Surgeons continued to teach giving crystalloid fluid for volume resuscitation to maintain blood pressure and red cells to maintain oxygen transport.

The first decade of this century found the US and other allied militaries in new wars in Afghanistan and Iraq. The large numbers of seriously injured patients presented new challenges for the military. Blood product support doctrine became controversial as traditional blood logistic assumptions conflicted with evolving surgical doctrine based on successfully treating the most severely injured. The contentions about appropriate blood supply and product use played out in the medical realm as arguments about (1) the acute coagulopathy of trauma, (2) the best way to resuscitate, (3) the best way to provide plasma and platelet coagulation support, and (4) the best way to get new products to the field. This paper will describe how progress in these four areas has changed field medical care in the last decade and address field blood use today.

The acute coagulopathy of traumaThe existence of an acute coagulopathy of trauma had been demonstrated in casualties in Vietnam7 and in animal models of soft tissue injury.8 Hematologists deemed it the early hemorrhagic phase of disseminated intravascular coagulation (DIC)9 explainable by the known concentrations and activities of the plasma coagulation factors.10 However, Brohi and colleagues in 2003 showed this to be a common clinical event, occurring in up to 25% of a thousand severely injured blunt trauma patients brought to the Royal London Hospital by helicopter before significant fluid administration. This observation led to a rethinking of the Advanced Trauma Life Support (ATLS) paradigm for trauma resuscitation in a few centers.11 Clearly, there were patients whose condition was likely being made worse by volume resuscitation with crystalloid, particularly when coupled with the use of plasma-poor packed red cells in additive solution. The most severely injured of these patients were bleeding and being transfused fast enough that by the time simple laboratory tests such as the prothrombin time (PT), partial thromboplastin time (PTT) and platelet count became available to guide therapy, the patients were profoundly coagulopathic and difficult to rescue with conventional blood products.

What Brohi and his colleagues specifically noted was that increases in the PT greater than 1.5 times normal in blood samples obtained at admission, became increasingly frequent as injury severity increased. Further, the proportion of patients with abnormal values increased from 10% among moderately injured patients to 80% among those with multiple profound injuries. When compared among patients with equivalent injury severity, those with a prolonged PT had four times greater mortality.

In a larger study published a month later, MacLeod and her colleagues in Miami examined the records of 20,103 patients admitted to the Ryder Trauma Center directly from the scene of injury.12 They found that any increase of the PT or PTT above normal was associated with excess mortality. In their study, increases in the PT were common, occurring in 28% of patients, but only 8% had an increased PTT. On the other hand, the odds ratios for death to be associated with an abnormal test were 3.6 for an increased PT and 7.8 for an elevated PTT. When adjusted for age and injury severity, the odds ratios were reduced to 1.35-fold excess mortality for an elevated PT and 4.26 for a prolonged PTT. The PT appeared to be a sensitive indicator of the acute coagulopathy of trauma, and the PTT was a specific marker of severe coagulopathy. The MacLeod group did not find an effect of admission platelet counts.


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Hess and colleagues went on and examined all 35,000 direct admissions to the University of Maryland Shock Trauma Center in Baltimore over a 7-year period.13 They demonstrated that increasing injury severity was associated with an increasing prevalence of prolongation of the PT and PTT and decreased concentrations of fibrinogen and platelets. In turn, increasingly abnormal values of these basic admitting laboratory tests were directly and strongly correlated with increased mortality.

The findings in these three series involving 56,000 patients are consistent with the animal models showing that serious soft tissue injury leads to a consumptive coagulopathy. The consumption is both direct, within the damaged tissue, and indirect through the thrombin interaction with thrombomodulin on remaining healthy cells leading to the activation of protein C and the inactivation of activated coagulation factors V and VIII and plasminogen activator inhibitor (PAI).14 The loss of PAI activity allows the unimpeded activation of plasmin and fibrinolysis. The most severely injured patients have increased PTs and fibrin breakdown products and reduced fibrinogen and platelet concentrations at levels that allow them to meet the International Society of Thrombosis and Hemostasis definition of DIC, but many other severely injured patients have subtler abnormalities that are still strongly associated with excess mortality.15 The work of the Maryland group showed that normal admission values of the PT and PTT that were in the upper half of the normal range and fibrinogen and platelets that were in the lower half of the normal range were also associated with excess mortality. In sum, these data affirmed the existence of an acute coagulopathy of trauma that occurs when injury severity exceeds individual capacity to compensate. Niles and her colleagues found the same patterns in combat wounded suffering penetrating injury.16 The most severely injured patients will arrive at sites of field medical care with coagulopathy already present.

Shock is often present in these severely injured patients. As a measure of physiologic injury, shock can be more important than calculated anatomic injury severity scores. Shock also serves as a mechanism exacerbating coagulation factor consumption by reducing blood flow and slowing thrombin clearance. As a result, the acute coagulopathy of trauma has also been called the acute coagulopathy of trauma and shock.

The best way to resuscitateFor thirty years, the American College of Surgeons’ Acute Trauma Life Support

course taught a generation of physicians that the proper way to resuscitate an injured hypotensive patient was to start two large bore IVs and infuse a 2 L bolus of crystalloid fluid. If blood pressure did not return to normal or if blood pressured did return to normal and subsequently fell again or if ongoing bleeding in excess of 100 mL/min was present, starting red cell transfusion to maintain oxygen carrying capacity was indicated, but crystalloid fluid was also given to maintain tissue perfusion. Laboratory tests were supposed to guide the administration of other blood components to keep the PT and PTT less than 1.5 normal, the fibrinogen greater than 1 g/L and the platelet count greater than 50 x 109/L. This plan of treatment assumed that coagulopathy is uncommon, develops late in care mostly as a result of dilution, and is relatively well tolerated. The new data suggested that for the most severely injured patients, none of these assumptions were true.

The ATLS resuscitation strategy violated classic teachings to limit resuscitation. In World War I, Cannon observed ‘‘If the pressure is raised before the surgeon is ready to check any bleeding that may take place, blood that is sorely needed may be lost”.17 In World War II, Beecher noted, ‘‘When the patient must wait for a considerable period, elevation of his systolic blood pressure to 85 mm Hg is all that is necessary.18 Balanced salt solutions were used in conjunction with immediate surgery in Vietnam,19 but calls for moderation in their use were largely unheeded.20 Efforts to reduce the volume of crystalloid fluids administered after the Vietnam War began with studies in animals with large vascular injuries. In swine with induced 4 mm aortic tears, resuscitation to normal blood pressures resulted in increased bleeding, coagulopathy, and mortality when compared to animals untreated and remaining hypotensive for a time.21 A randomized human trial of permissive hypotension following penetrating truncal injury published by Bickell and colleagues as a lead article in The New England Journal of Medicine in 1994 showed an 8% lower mortality in the patients not given fluids before surgery.5 For moderately injured patients, improved outcomes were noted with permissive hypotension, rapid diagnosis of bleeding location and early surgery.

For the most severely injured, death from uncontrolled hemorrhage can occur within minutes of injury, many dying before medical care arrives on the scene. Nevertheless, urban and helicopter ambulance systems delivered such patients alive to academic Level 1 trauma centers with increasing frequency. A number of trauma centers, including Helsinki, Sydney, Houston, Seattle, Denver, and Baltimore adopted systems of resuscitating the most severely injured with blood


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components, using red cells, plasma, and platelets in 1:1:1 ratios.22 In Baltimore, a retrospective review of a year of blood usage in the Shock Trauma Center by Como and his colleagues, demonstrated the 1:1:1 ratio, but also that the red cells tended to be given early and the plasma and platelets late, in an attempt to rescue coagulopathic patients.23

By 2004, the rate and severity of combat casualties seen in the combat support hospitals in Iraq had worsened. Resuscitation with crystalloids and universal donor red cells frequently led to greater than ten unit transfusions and death from uncontrolled hemorrhage before blood could be typed and plasma thawed. Clinicians decided to transfuse equal numbers of units of plasma and RBC, while minimizing crystalloids. Results appeared to improve.24 However, the surgeons who lived the experience, remember the rapid control of hemorrhage with plasma use, as well as being able to close wounds primarily and get patients off of ventilators quickly when edema from crystalloid fluid was minimized.

Academic arguments about plasma use in resuscitation tend to focus on the question of whether 1:1:1 is the right blood product ratio, a question whose answer is likely to be highly dependent on the injury severity and rate of bleeding of the population under study. Cotton and colleagues have shown that 1:1:1 resuscitation is associated with a decrease in the fraction of all patients transfused being massively transfused, reduced total blood use, and lower mortality among transfused patients.25 Kautza and colleagues confirmed the decreasing frequency of massive transfusion in a large multi-institutional cohort receiving more plasma and platelets over the period 2004-2009.26 A prospective observational study in ten US trauma centers showed that more than 2.5 h were required to deliver plasma and platelets to critically injured, bleeding, and red cell-transfused patients.27 A multicenter prospective randomized clinical trial is in progress.28

How to deliver plasma and platelet support in the fieldLarge civilian hospitals and medical centers in developed countries supply their needs for therapeutic plasma with thawed frozen or fresh frozen plasma (FFP) or, rarely, liquid plasma and for platelets with five-to-seven-unit pools of whole-blood-derived platelet concentrates or single-donor apheresis platelets. The products are readily available from local donor centers and additional components needed in times of high demand are available in national blood systems. These standard products are safe and effective within our understanding

of the terms, but neither is easy to support in field medical care. In fact, the short storage duration of thawed plasma and platelets makes their use inefficient in small general hospitals and has led to these critical products being unavailable when seriously bleeding patients are seen in smaller or remote facilities.

PlasmaPlasma is generally stored and transported frozen because, once thawed, it must be used within 5 days or discarded. The current generation of polyvinyl chloride bags is clear, tough, and malleable when thawed and make plasma easy to inspect for quality control and administer. However, the bags become brittle at the temperatures of the “dry ice” that are used in shipping.29 Bag breakage rates as high as 50% have been reported, and rates of 20% are widely accepted as normal. Once delivered to a remote location, plasma needs to stay frozen until shortly before use which means that electricity and freezers need to be available constantly. On the other hand, at the time of use, up to 30 min may be required to thaw a unit of ABO compatible plasma by immersion in 37°C water. Bag fractures, detected only at the time of thawing, reduce the efficiency of field use, contaminate the thawing system, and cleanup can delay the process of issue further.

Plasma can be provided in alternate forms. Freeze-dried plasma was widely used by the US military during World War II.30 Several countries make freeze-dried plasma for military and remote use including Germany, Belgium, France and Australia. Collecting universal donor AB plasma by apheresis in 600 mL amounts from single donors, rotary freezing of the anticoagulated plasma in bottles, and cold vacuum drying is the easiest method that meets international standards, but the process does not scale easily and has risks of cross-contamination. Units also can be made from pools of whole plasma from many donors followed by pathogen reduction such as solvent/detergent treatment and nanofiltration and are compatible with industrial lot quality control and mass manufacture. However, industrially processed plasma from pools suffers the consequences of the additional processing in the loss of labile factors. The original such product, produced for a short time in the US, was associated with thrombotic episodes thought to be related to low concentrations of protein S.31 As a result, no such product was available in the US until 2013. A disadvantage of the whole approach of providing unfractionated plasma for resuscitation is the variable but low concentration of coagulation factors in any unfractionated


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plasma product such as FFP. Most of the protein present in such products is albumin, which because of its strong colloid osmotic activity, leads to dilution of the coagulation proteins to their original concentrations in the donor plasma or even lower because of losses in processing.32

An alternative approach to delivering plasma coagulation factors is to separate the plasma protein components and prepare them as pathogen-reduced concentrated products. This has been performed with fibrinogen, anti-hemophilia factors, and vitamin K dependent factors for many decades. The resulting products are well-defined, widely used in medical care, and moderately to quite expensive. European groups have pointed out that the combination of fibrinogen concentrates and 4-factor prothrombin complex concentrates (PCCs) can reconstitute the extrinsic coagulation pathway when these materials are used with platelets as a source of factor V. This works because endothelium-derived factor VIII is rarely reduced and factor XIII requirements are met by the amounts in the fibrinogen concentrates. Schochl and his colleagues at the Vienna trauma center have reported that patients resuscitated with fibrinogen and PCCs have better outcomes and require fewer blood components than patients resuscitated with conventional blood products.33,34 Larger, albeit still uncontrolled, experience supporting this concept, has been reported.35-37

PlateletsProviding platelets in the field has been a major challenge. Liquid stored platelets have a 5-day shelf life and storage requirements that can only be met in heated or airconditioned buildings. With conventional collection and processing followed by direct air delivery, they have only a 3-day shelf life remaining and then require a mechanical agitator in an environmental box capable of maintaining a temperature between 20 and 24°C. Frozen platelets can be made from single donor collections processed into 4-10% dimethyl sulfoxide (DMSO) and frozen in mechanical freezers or liquid nitrogen.38,39 Prior to freezing, the products can be reduced to very small volumes (around 10 mL) and then kept for at least 3 years.40 DMSO frozen platelets have been transfused successfully since the 1970s.41 Despite 50% lower functional recovery than that of fresh liquid platelets, frozen platelets are more effective in shortening the skin bleeding time compared to 3-day liquid-stored platelets.42 Frozen platelets also show a higher capacity to bind factor V and a higher thromboxane production after ADP stimulation. The Dutch military used them extensively in Iraq and Afghanistan and so did the

Australians, who depended on the Dutch blood supply in Afghanistan.43,44 The results reported were excellent, but at this point the Dutch are the only group in the world with extensive experience with them.

A variety of platelet substitutes or platelet extenders have been proposed including fibrinogen-coated albumin microspheres, thromboerythrocytes, and stabilized platelet membrane fragments. Each of these substitutes restores some but not all platelet functions, require extensive manufacturing and clinical validation, and lack clot-retraction potential. As a result, frozen platelets remain the most attractive of these products in the short term. Frozen platelets have relative ease of manufacture, long and stable storage, ease of use in a thaw and infuse mode, safety from allergic and immune reactions due to their markedly reduced plasma content and protection from bacterial growth, a history of successful clinical use, and the ability to respond to variable demand. All of these factors suggest that frozen platelets will translate into conventional blood banking. The reduced losses of liquid platelets to outdating associated with using a mixed inventory of liquid and frozen platelets would help offset the higher cost of the frozen product. In turn, a large civilian inventory of frozen plate-lets would mean that the products would be available in times of disaster or war. Military use is generally a very small fraction of civilian use but with highly variable use rates. This pattern of episodic urgent demand makes longer storage a critical feature. Small thermocouple cascade (-80°C) freezers to support their use in the field are already available.

Getting new blood products to the fieldAttempts to develop and license new blood storage technologies for military and austere medical use have been limited both by the low rate of licensure of all blood products and the extremely limited market for blood products usable in the field. This has left the military as essentially the sole funder of the effort to develop novel blood systems. Luckily, the militaries of a number of developed countries have active programs and have championed widely different products. The Dutch development of frozen platelets for use in Iraq and Afghanistan and the Australian work with freeze-dried plasma after their experiences in East Timor are examples. US Special Operations Forces medics carry French-manufactured freeze-dried plasma in glass bottles in prehospital field operations in Afghanistan.


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The US military has at once the largest blood product development programs and the most difficult regulatory barriers to fielding new products. The US military has active programs in prolonged red blood cell storage, frozen platelets, and freeze-dried plasma. The US Food and Drug Administration (FDA) enforces requirements that such new blood products be proven safe and effective before licensure. However, as definitions of effectiveness move from the historic surrogate measures of increased concentrations of the transfused product to large clinical studies, the time and cost of developing new products has grown. The end result is that products developed by the US military such as 8-week red cell storage and frozen platelets are licensed in Europe, but not in the US. The developmental times for these products has been more than two decades, creating problems in scientific developmental and regulatory continuity.

The Dutch system is more efficient with their military blood bank integrated into their national blood system. Novel products are made, tested, and approved for use within the system, which has national responsibility for blood provision and safety. In a small country where almost most all of the expertise in blood banking is in the national blood banking system, self-regulation makes sense.

The heart of the problem of providing blood support for field medical care is envisioning a set of blood products that can work in the field and at small regional hospitals. Long life liquid RBCs, pharmaceutically manufactured fibrinogen and other coagulation factor concentrates, dried plasma, and small-volume frozen platelets all share a potential for wide use in conventional clinical medicine leading to large standing inventories, ease of transport, availability for rapid use, and compatibility with the field environment. Items that work well in the field fit on the pallet with other emergency medical supplies as a surgical team heads to a disaster and are items that the doctors and nurses have seen and used before. Scientists, developers, and program managers need to commit to a small list of such products and gather the resources to get the products licensed and into practice.

Solving this conundrum of field blood product availability will also directly address the problem that half of trauma patients in the US are initially cared for in non-Level 1 trauma centers. This is a significant unmet national blood need. In the Maryland state trauma system, two Level 1 trauma centers can provide thawed plasma and readily available liquid platelets, three Level 2 centers have frozen plasma and access to platelets with a several hour delay and four Level 3 centers have frozen plasma but no rapid access to platelets. This pattern

is repeated in the 10 hospitals of the Memorial Hermann Hospital System in Houston, Texas. Nationally, smaller trauma centers have adequate numbers of liquid RBCs, only limited amounts of frozen plasma, no thawed plasma, and, frequently, no platelets at all. They are forced by this pattern of product availability to continue transfusion patterns that did not work in the wars. Dried plasma and frozen, low-volume platelets can provide the benefits of hemorrhage control resuscitation for many more citizens. PCCs and fibrinogen concentrates offer a potential alternate route to providing plasma rapidly, and have the advantage of being commercially available now, but await randomized trials to demonstrate at least non-inferiority. Such trials would also highlight economic and logistic differences between the products.

Addressing field blood use todayIn practice in the war zones, each country has its own national mix of products. The Dutch have a number of liquid-frozen blood bank modules, fielding liquid red cells, frozen red cells, frozen plasma, and frozen platelets. A bilateral agreement on mutual support between the US and Netherlands blood programs proved very effective, especially in Iraq. Under this agreement, the Dutch got liquid red cells from the Americans when they needed RBCs faster than frozen RBCs could be thawed and deglycerolized, and the Americans got frozen platelets from the Dutch, because they had these products available for emergencies. As the current conflicts wind down, blood product use will decrease, and the shared products become less available. The reliance on fresh whole blood as the backup to the currently inadequate US product mix will remain.

Surgical teams deploying to civil disasters such as the earthquakes in Turkey, Iran, Pakistan, Haiti, and China need to have the entire spectrum of blood support plans.45 Most blood will be used early but some will be required for secondary operations. An Israeli surgical team, sent to the Turkish earthquake, took twenty units of red cells and used five. Iran and China had national blood systems which responded appropriately to national disasters in the last decade. Haiti’s blood system was inadequate before the earthquake and was destroyed. Teams need to plan for all eventualities. Potential donors of universal donor blood among team members need to be identified in advance and pretested before deployment. Donation, storage, and documentation need to be preplanned. At this time, transfusion of whole blood should be planned and a system to support its use developed.


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Transfusion of RBCs is not the same as transfusing blood. RBC transfusion remains an adjunct to resuscitative surgical care, only occasionally is it lifesaving by itself. Optimal transfusion of other blood components is important, but required amounts remain in question. Balancing the real albeit small risk of transfusion with the significant potential benefit remains a challenge. Current clinical data suggest early transfusion of balanced amounts of blood products decreases overall blood product use, minimizes morbidity and improves outcome. Transitioning this experience to remote facilities, whether they are in military theaters or small civilian hospitals remains a challenge. Working with regulators to develop a set of products that comply with safety rules and yet are logistically feasible and clinically effective across the entire spectrum of transfusion locations is the goal. We think this goal is obtainable.

References

1. Holcomb JB, Spinella PC. Optimal use of blood in trauma patients. Biologicals 2010;38(1):72-7.

2. Hess JR, Thomas MJG. Blood use in war and disaster: lessons from the past century. Transfusion 2003;43(11):1622-33.

3. Rotondo MF, Schwab CW, McGonigal MD, Phillips GR, Fruchterman TM, Kauder DR, Latenser BA, Angood PA. ‘Damage control’: an approach for improved survival in exsanguinating penetrating abdominal injury. J.Trauma 1993;35(3):375-82.

4. Thorson CM, Dubose JJ, Rhee P, Knuth TE, Dorlac WC, Bailey JA, Garcia GD, Ryan ML, Van Haren RM, Proctor KG. Military trauma training at civilian centers: a decade of advancements. J.Trauma Acute.Care Surg. 2012;73(6 Suppl 5):S483-S489.

5. Bickell WH, Wall MJJ, Pepe PE, Martin RR, Ginger VF, Allen MK, Mattox KL. Immediate versus delayed fluid resuscitation for hypotensive patients with penetrating torso injuries. N.Engl.J.Med 1994;331(17):1105-9.

6. Dutton RP, Mackenzie CF, Scalea TM. Hypotensive resuscitation during active hemorrhage: impact on in-hospital mortality. J.Trauma 2002;52(6):1141-6.

7. Simmons RL, Collins JA, Heisterkamp CA, Mills DE, Andren R, Phillips LL. Coagulation disorders in combat casualties. I. Acute changes after wounding. II. Effects of massive transfusion. 3. Post-resuscitative changes. Ann.Surg. 1969;169(4):455-82.

8. Hardaway RM. Shock and disseminated intravascular coagulation. Thromb.Diath.Haemorrh.Suppl 1966;20:121-46.

9. Levi M, Ten Cate H. Disseminated intravascular coagulation. N.Engl.J.Med 1999;341(8):586-92.

10. Armand R, Hess JR. Treating coagulopathy in trauma patients. Transfus.Med Rev. 2003;17(3):223-31.

11. Brohi K, Singh J, Heron M, Coats T. Acute traumatic coagulopathy. J.Trauma 2003;54(6):1127-30.

12. MacLeod JBA, Lynn M, McKenney MG, Cohn SM, Murtha M. Early coagulopathy predicts mortality in trauma. J.Trauma 2003;55(1):39-44.

13. Hess JR, Lindell AL, Stansbury LG, Dutton RP, Scalea TM. The prevalence of abnormal results of conventional coagulation tests on admission to a trauma center. Transfusion 2009;49(1):34-9.

14. Hess JR, Brohi K, Dutton RP, Hauser CJ, Holcomb JB, Kluger Y, Mackway-Jones K, Parr MJ, Rizoli SB, Yukioka T, et al. The coagulopathy of trauma: a review of mechanisms. J.Trauma 2008;65(4):748-54.

15. Taylor FBJ, Toh CH, Hoots WK, Wada H, Levi M. Towards definition, clinical and laboratory criteria, and a scoring system for disseminated intravascular coagulation. Thromb.Haemost. 2001;86(5):1327-30.


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16. Niles SE, McLaughlin DF, Perkins JG, Wade CE, Li Y, Spinella PC, Holcomb JB. Increased mortality associated with the early coagulopathy of trauma in combat casualties. J.Trauma 2008;64(6):1459-63.

17. Cannon WB FJCE. The preventive treatment of wound shock. J Am Med Assoc 1918;70:618-21.

18. Beecher HK. Preparation of Battle Casualties for Surgery. Ann.Surg. 1945;121(6):769-92.

19. Carrico CJ, Canizaro PC, Shires GT. Fluid resuscitation following injury: rationale for the use of balanced salt solutions. Crit Care Med 1976;4(2):46-54.

20. Moore FD, Shires GT. Moderation. Anesth.Analg. 1968;47(5):506-8.21. Bickell WH, Bruttig SP, Millnamow GA, O’Benar J, Wade CE. The detrimental effects

of intravenous crystalloid after aortotomy in swine. Surgery 1991;110(3):529-36.22. Malone DL, Hess JR, Fingerhut A. Massive transfusion practices around the globe

and a suggestion for a common massive transfusion protocol. J.Trauma 2006;60(6 Suppl):S91-S96.

23. Como JJ, Dutton RP, Scalea TM, Edelman BB, Hess JR. Blood transfusion rates in the care of acute trauma. Transfusion 2004;44(6):809-13.

24. Borgman MA, Spinella PC, Perkins JG, Grathwohl KW, Repine T, Beekley AC, Sebesta J, Jenkins D, Wade CE, Holcomb JB. The ratio of blood products transfused affects mortality in patients receiving massive transfusions at a combat support hospital. J.Trauma 2007;63(4):805-13.

25. Cotton BA, Gunter OL, Isbell J, Au BK, Robertson AM, Morris JAJ, St Jacques P, Young PP. Damage control hematology: the impact of a trauma exsanguination protocol on survival and blood product utilization. J.Trauma 2008;64(5):1177-82.

26. Kautza BC, Cohen MJ, Cuschieri J, Minei JP, Brackenridge SC, Maier RV, Harbrecht BG, Moore EE, Billiar TR, Peitzman AB, et al. Changes in massive transfusion over time: an early shift in the right direction? J.Trauma Acute.Care Surg. 2012;72(1):106-11.

27. Holcomb JB. For the PROMMTT study group. The prospective, observational, multicenter, major trauma transfusion (PROMMTT) study: comparative effectiveness of a time-varying treatment with competing risks. Arch Surg 2012;15:1-10.

28. Clinical Trials.gov Identifier NCT01545232. 2012.29. Hmel PJ, Kennedy A, Quiles JG, Gorogias M, Seelbaugh JP, Morrissette CR, Van

Ness K, Reid TJ. Physical and thermal properties of blood storage bags: implications for shipping frozen components on dry ice. Transfusion 2002;42(7):836-46.

30. Kendrick DB. Blood Program in World War II. Washington DC: Office of the Surgeon General; 1964.

31. Flamholz R, Jeon HR, Baron JM, Baron BW. Study of three patients with thrombotic thrombocytopenic purpura exchanged with solvent/detergent-treated plasma: is its decreased protein S activity clinically related to their development of deep venous thromboses? J.Clin.Apher. 2000;15(3):169-72.

32. Solheim BG, Seghatchian J. Update on pathogen reduction technology for therapeutic plasma: an overview. Transfus.Apher.Sci. 2006;35(1):83-90.

33. Schochl H, Nienaber U, Hofer G, Voelckel W, Jambor C, Scharbert G, Kozek-Langenecker S, Solomon C. Goal-directed coagulation management of major trauma patients using thromboelastometry (ROTEM)-guided administration of fibrinogen concentrate and prothrombin complex concentrate. Crit Care 2010;14(2):R55.

34. Gorlinger K, Fries D, Dirkmann D, Weber C, Hanke A, Schochl H. Reduction of Fresh Frozen Plasma Requirements by Perioperative Point-of-Care Coagulation Management with Early Calculated Goal-Directed Therapy. Transfus.Med Hemother. 2012;39(2):104-13.

35. Joseph B, Amini A, Friese RS, Houdek M, Hays D, Kulvatunyou N, Wynne J, O’Keeffe T, Latifi R, Rhee P. Factor IX complex for the correction of traumatic coagulopathy. J.Trauma Acute.Care Surg. 2012;72(4):828-34.

36. Innerhofer P, Westermann I, Tauber H, Breitkopf R, Fries D, Kastenberger T, El Attal R, Strasak A, Mittermayr M. The exclusive use of coagulation factor concentrates enables reversal of coagulopathy and decreases transfusion rates in patients with major blunt trauma. Injury 2013;44(2):209-16.

37. McSwain Jr N BJ. Potential use of prothrombin complex concentrate in trauma resuscitation. J Trauma 2011; 70(5 Suppl.):S53-6 2011.

38. Lazarus HM, Kaniecki-Green EA, Warm SE, Aikawa M, Herzig RH. Therapeutic effectiveness of frozen platelet concentrates for transfusion. Blood 1981;57(2):243-9.


40. Schiffer CA, Aisner J, Wiernik PH. Clinical experience with transfusion of cryopreserved platelets. Br.J.Haematol. 1976;34(3):377-85.

41. Daly PA, Schiffer CA, Aisner J, Wiernik PH. Successful transfusion of platelets cryopreserved for more than 3 years. Blood 1979;54(5):1023-7.



44. Neuhaus SJ, Wishaw K, Lelkens C. Australian experience with frozen blood products on military operations. Med J Aust. 2010;192(4):203-5.

45. Abolghasemi H, Radfar MH, Tabatabaee M, Hosseini-Divkolayee NS, Burkle FMJ. Revisiting blood transfusion preparedness: experience from the Bam earthquake response. Prehosp.Disaster.Med 2008;23(5):391-4.

Chapter

General discussion

8


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BackgroundBased on article 97 of the Constitution of the Kingdom of the Netherlands, the armed forces “shall not only defend and protect the interests of the Kingdom, but also maintain and promote the international legal order”. During the past decade, this has been mainly reflected in the large scale (military) participation of the Netherlands in missions in Iraq and Afghanistan. The common denominators of such deployments are the large distances between home country and area of operations, as well as the frequently high intensity of the conflicts, with sometimes considerable numbers of heavily injured, massively bleeding casualties. Despite technological developments over the past decades, exsanguination because of massive hemorrhage still remains the major cause of death on the battlefield.1-6

Damage Control ResuscitationToday’s treatment of massively injured patients follows the principles of damage control resuscitation (DCR).7 It incorporates two elements that are integral to each other and never should be practiced separately8: damage control surgery9 and (massive) blood transfusion. The former mainly consists of a quick, short surgical intervention to control life-threatening hemorrhages and peritoneal contamination by intraperitoneal packing and temporary closure. Re-exploration and definitive surgery are postponed until the patient’s physiology has been sufficiently restored. The latter, blood transfusion, is aimed at preventing or correcting the so-called “bloody vicious cycle”, the lethal triad of hypothermia, acidosis and coagulation defects.10

Battlefield damage control resuscitationIn a battle zone, the first and most forward surgical capacity is present at the level of Forward Surgical Teams (FST) or equivalents thereof. Regardless of exact size and number of personnel, their most important task is to perform the necessary life-saving surgical interventions on non-compressible and truncal bleeds, if any way possible within the “golden hour” of injury. Adhering to the principles of DCR, it is absolutely necessary to enable blood transfusion at FST-level as an indispensable adjunct to restore blood volume, oxygen carrying capacity and correct clotting defects. This means that the capacity of an FST, deployed with the primary intent to save lives as early as possible after injury, is (partly) wasted, if it does not have the (sufficient) capacity to perform the necessary blood transfusions to complement the DCR. On the other hand, transfusing (whole)

blood or blood components into a profusely bleeding patient is useless without having surgical capacity available to explore and treat non-compressible bleeds.The logistical challenge includes supplying the surgical treatment facilities in theater with enough blood products to treat unknown numbers of hemorrhaging casualties at unpredictable moments. While maintaining viability, it implies at least storage of blood products, as close as possible to the point of care. These aspects were for the first time addressed and documented during World War I, when whole blood was successfully stored.11-13

Whole blood and blood componentsThe development of blood component therapy in the 1960s and 70s14 changed the standard transfusion practice, not only in the civilian setting, but also on the battlefield. It enabled a more efficient use of donated whole blood by limiting unnecessary transfusions and avoiding wastage due to outdating, with optimal storage conditions for each component. Worldwide, transfusion of whole blood has thus substantially decreased, but (mainly) US military experiences from Iraq and Afghanistan over the past decade have revived interest in this approach,15-21 especially in situations of massive transfusions, even without the availability of surgical capacity.22-26

The rationale for this approach has primarily been the fact that military blood logistics, relying on standard blood components, is sometimes unable to keep up with the needs of the trauma surgeons in the field, particularly in the case of massive transfusions in forward surgical settings. Whole blood, derived from on-site personnel in theater, is then used as a supplement to or even replacement of standard blood components. It has to be acknowledged that fresh (warm) whole blood is capable of correcting hypothermia, as well as serious deficits in the coagulation system, circulating volume and oxygen carrying capacity. However, despite claimed advantages such as lower donor exposure, the natural, right ratio between constituents, higher hematocrit, clotting factor content and platelet count,15,16 there are undeniable downsides. In order to effectuate the use of (fresh) whole blood, one needs to set up and maintain a system with all suitable donors on site (“walking blood bank”), while at the same time keeping track of people rotating in and out. This is a major logistic effort, involving lots of time and personnel: not only pre-deployment immunization and screening, but also reliable, approved pre-donation (rapid) testing for transfusion transmitted diseases (TTDs) should be incorporated. Cases of Hepatitis B, C 25,27 and Acute


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Lung Injury (ALI) 28 have been reported in conjunction with transfusing whole blood, as well as HTLV 29 transmission and even (fatal) transfusion associated graft-versus-host disease.30 Furthermore, donor testing in theater should be repeated at regular intervals.31,32 If not only group O (low titer anti-A, anti-B) whole blood is used,33,34 precious time will also be lost to perform a crossmatch. If the current international standards of leukodepletion would be followed, while attempting to save the platelets in a whole blood unit, the application of whole blood platelet-sparing filters would add more than 30 min to the processing of donated blood.35 Finally, although whole blood, if stored in CPDA-1 at 2-6⁰C, expires after 35 days,36 it retains its hemostatic properties for up to only 21 days 20 and its platelet function for only 15 days.37

The use of (fresh) whole blood, even on the battlefield, remains therefore controversial. Up till now, no study has ever indisputably demonstrated the clinical superiority of whole blood over the triplet of essential components red cells, plasma and platelets, let alone over the frozen equivalent of these blood components. To date, whole blood, however, is definitely less safe than fully tested components from fully tested, regular donors. All in all, the use of whole blood, even on the battlefield, should be made redundant, by putting the necessary efforts in exploring other options to meet the transfusion needs of a massively bleeding patient. What is needed, therefore, is a way to solve the logistic issues of today’s high intensity, far-off conflicts, while at the same time achieving the beneficiary effects of rapidly available blood transfusion, without exposing recipients to unnecessary risks. Longer shelf lives of fully tested red cells (and platelets), while retaining their essential functions, would help remedy the problems of time and distance factors.

Extending the current shelf life of (liquid) red blood cells, measured in weeks, to years is only possible at sub-zero temperatures (cryopreservation). This poses an obstacle, because freezing red cells without any further precautions usually causes irreparable and lethal damage, due to increasing hypertonicity and ice crystal formation.

Cryopreservation The most practical approach to prevent ice formation and the accompanying damage is to use cryoprotectants, compounds that are capable of reducing damage to cells during freezing. After the accidental discovery of glycerol as a useful cryoprotectant in preserving cell viability,38 it became rapidly apparent39

that red cells were also excellently preserved at relatively high temperatures. Since then, a variety of freezing techniques has been developed, modified and used.40-46

Methods of cryopreservation The ideal of a one-step procedure, requiring no additional processing, is still far from reality, be it ever achieved. Only two methods survived for clinical use, both using glycerol as a cryoprotectant in the two-step technique.

1. The low-glycerol / rapid freezing technique2. The high-glycerol / slow freezing technique

The low-glycerol method (LGM) allows the use of low concentrations of glycerol (17-20%), but requires rapid freezing (>100°C/min) and storage at temperatures below -150°C (vapor phase of liquid nitrogen). The reasons for developing this technique was the idea that lower concentrations would cause less osmotic stress, enable shorter postthaw processing time and at the same time an almost infinite storage period, because below -130°C cell metabolism comes to a complete stand-still.47,48 The mandatory use of liquid nitrogen, however, simply precludes this method from being considered suitable for military, operational purposes.

The high-glycerol method (HGM) requires higher concentrations (40-50%) of glycerol, therefore a longer postthaw processing time, but enables slower freezing rates (1-3°C/min) and higher storage temperatures of below -65°C, but usually around -80°C. This means that mechanical freezers can be used for storage. Although cell metabolism at these temperatures is very much depressed, but not completely halted, storage times exceeding 30 years are possible.49 Within Europe only the UK has approved a storage time of 30 years for frozen red cells.50 The wording of current European and US guidelines,36,51 however, leaves room for further extension, in accordance with the UK.

Deglycerolization (ACP215)Regardless of the glycerol concentration used, all units need to be deglycerolized after thawing, to reduce the glycerol concentration to around 1%, in order to avoid hemolysis in the recipient’s circulation.52 The introduction in 1998 of the functionally closed, automated cell processor ACP215 (Haemonetics®) meant a


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major step forward in the applicability of frozen red cells daily practice, because it enabled postthaw storage for at least two weeks in AS-3 at 2-6°C.53 Until then, all techniques used to remove the glycerol were considered “open”, meaning a possibility of bacterial contamination. Thus, red cells once thawed and washed, needed to be transfused within 24 hr.

Obviously, the equipment needed to process frozen red cells, substantially adds to the costs of these blood products, as compared to standard liquid red cells. Nonetheless, frozen red cells have several advantages over stored standard liquid red cells, paradoxically mainly related to the mandatory deglycerolization procedure. Actually, deglycerolization not only removes the cryoprotectant, but also eliminates otherwise unwanted substances like cell debris, cytokines and free Hb.54 By consequence, patients are treated with a “clean” product, if transfused immediately after the washing procedure.

In chapter 2 we compared the LGM and HGM, using the ACP215. HGM-red cells showed better stability during storage in SAGM at 2-6°C after deglycerolization.55 Also, postthaw storage in AS-3 demonstrated reduced hemolysis, compared to SAGM, as was already shown in earlier studies.56,57 These results were the reference points for further studies, aimed at improving the practical applicability of frozen red cells during worldwide military missions under austere conditions.

Chapters 3 and 4 discussed the experiences with the HGM during deployments in Southwest Asia from a Dutch and Australian perspective.58,59 These two reports showed that frozen blood products, including red cells, despite higher cost, not only substantially reduce the number of resupply shipments, but also enable the abolition of the “walking blood bank”, without compromising the in-theater availability of safe and effective blood products, all in compliance with international regulations and guidelines.36,51

Adjustments in the HGM procedureIt does not seem likely that the currently used deglycerolization device will be improved in the sense that its processing time will be considerably shortened. Since time is of the utmost essence in treating massive blood loss patients, the possibilities to improve the practical applications of frozen red cells therefore mainly should be found in adapting the procedure around the use of the device, to gain extra postthaw storage time, without compromising the requirements of

international guidelines. This encompasses aspects like prefreeze interventions, the washing procedure and the postthaw storage conditions.

The first aspect was investigated and discussed in chapter 5 and encompassed a modified method of glycerolization, eliminating the centrifugation step that reduces the glycerolized RBC supernatant.60 It thus shortens the glycerolization time, but more importantly, the results confirmed the suggestion from List et al.61 that skipping the prefreeze centrifugation step leads to a better postthaw shelf life of previously frozen red cells at 2-6°C. Most probably, skipping the centrifugation step in the glycerolization procedure plays an important role in minimizing shear stress on the red cells,62,63 resulting in less damage, and thus better storable thawed red cells.

The second aspect encompasses a modification in the washing procedure by using phosphate buffered saline (PBS) with a higher pH (7.4) than the normally used 0.2% glucose/0.9% NaCl-mixture as described before.64 The use of PBS helped to maintain ATP at higher levels during deglycerolization. While maintaining the standard AS-3 final resuspension medium, the two previous adjustments prolonged postthaw storage shelf life to 28 days at 2-6°C.

In chapter 6 we addressed a third aspect, prefreeze biochemical intervention, called rejuvenation, to raise the red cell’s ATP, total adenylate and 2,3 DPG content to normal or even supranormal levels.65 Rejuvenation will also, at least partially, restore the shape change from spherocytes and echinocytes back to discs,66 improve O2-delivery,67 as well as the membrane’s elasticity68 and reverse post-storage RBC adhesion to endothelial cells.69,70

Total adenylate content in prefreeze Rejuvesol- and PAGGGM-treated red cells was maintained way above 82 % of the original values for at least 42 days, if they were stored in AS-3 after thawing and washing. Based on a very good correlation between total adenylate levels and “in vivo” survival,71 these results suggest a 24hr survival of >75%, as required by European and US guidelines.36,51 With regard to hemolysis at the time of infusion, a postthaw storage time of 35 days and 42 days would be allowed in Europe (0.8%) and the US (1.0%) respectively.36,51 In addition, the beneficial effect on 2,3-DPG levels after thawing and deglycerolization is maintained up to 14 days of storage.

The important practical implication of the above-mentioned studies is that it facilitates operating a frozen blood bank, by minimizing the effects of a time consuming deglycerolization procedure. One hour of deglycerolization per unit has less effect on the direct availability of red blood cells for transfusion, if each


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deglycerolized unit can be stored afterwards for at least 35 days, as compared to the current 14 days.36,51,72 Thus, the ability to maintain a liquid, previously frozen, inventory of red cells with an extended shelf life of 35 days creates the possibility to better meet unexpected operational demands, as is the case under battlefield conditions.

Chapter 7 reviewed the international advances in military transfusion medicine in the last decade, based on experiences of twenty years of war in Southwest Asia.73 The wars in Iraq and Afghanistan have highlighted the essential role of primary resuscitation with blood products in the care of critically injured soldiers. It stresses the problems of logistics not only in a military environment, but also in smaller civilian trauma centers. The common denominator is that both health care systems need help in assuring the immediate availability of sufficient blood products in case of emergencies. This implies the use of blood products with low logistic burdens, including longer shelf lives. Although not ideal, frozen blood products with longer postthaw shelf lives tick some of the most essential boxes.

Conclusion

Modifications of several steps in the freezing procedure of red cells offer practical solutions to enable ready availability of this essential blood component in the treatment of severe blood loss, even in wartime conditions. In addition to earlier studies that confirmed the safety and effectiveness of frozen red cells, 74-77 the application of these adjustments enable postthaw storage of red cells in AS-3 at 2-6°C for 35 days in Europe and 42 days in the US, taking into consideration both hemolysis at the time of infusion and 24 hr posttransfusion survival of > 75%.36,51,78 This extension to the equivalent of the standard red cell concentrates in civilian practice proves that an efficient use of frozen red cells is possible, even during military deployments.

Contrary to Hess,79 who predicted a small effect of red cell freezing on the logistics of the blood supply, the introduction of frozen red cells has had, at least for the Netherlands, a great impact on the military supply chain, particularly since the turn of the century. Frozen red cells, together with frozen plasma and frozen platelets, have become an important asset in treating hemorrhagic patients on the battlefield, as was recently reported by Noorman et al.80 who

carried out an in-depth analysis of using predominantly frozen blood products in battle casualties, including dozens of massive transfusions, with survival rates comparable to those in the civilian setting. Therefore, frozen blood products are definitely able to make the in-theater emergency whole blood collections redundant.


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References

1. Maughon JS. An inquiry into the nature of wounds resulting in killed in action in Vietnam. Mil.Med. 1970;135(1):8-13.

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11. Rous P, Turner JR. The preservation of living red blood cells in vitro: I. Methods of preservation. J.Exp.Med. 1916;23(2):219-37.

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17. Spinella PC, Perkins JG, Grathwohl KW, Repine T, Beekley AC, Sebesta J, Jenkins D, Azarow K, Holcomb JB. Risks associated with fresh whole blood and red blood cell transfusions in a combat support hospital. Crit Care Med 2007;35(11):2576-81.

18. Spinella PC. Warm fresh whole blood transfusion for severe hemorrhage: U.S. military and potential civilian applications. Crit Care Med 2008;36(7 Suppl):S340-S345.

19. Pidcoke HF, McFaul SJ, Ramasubramanian AK, Parida BK, Mora AG, Fedyk CG, Valdez-Delgado KK, Montgomery RK, Reddoch KM, Rodriguez AC, et al. Primary hemostatic capacity of whole blood: a comprehensive analysis of pathogen reduction and refrigeration effects over time. Transfusion 2013;53 Suppl 1:137S-49S.

20. Murdock AD, Berseus O, Hervig T, Strandenes G, Lunde TH. Whole blood: the future of traumatic hemorrhagic shock resuscitation. Shock 2014;41 Suppl 1:62-9.

21. Chandler MH, Roberts M, Sawyer M, Myers G. The US military experience with fresh whole blood during the conflicts in Iraq and Afghanistan. Semin.Cardiothorac.Vasc.Anesth. 2012;16(3):153-9.

22. Bowling F, Pennardt A. The use of fresh whole blood transfusions by the SOF medic for hemostatic resuscitation in the austere environment. J.Spec.Oper.Med 2010;10(3):25-35.

23. Bowling F, Kerr W. Fresh Whole Blood transfusions in the austere environment. J.Spec.Oper.Med 2011;11(3):3-37.

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25. Beckett A, Callum J, da Luz LT, Schmid J, Funk C, Glassberg E, Tien H. Fresh whole blood transfusion capability for Special Operations Forces. Can.J.Surg. 2015;58(3 Suppl 3):S153-S156.

26. Cordova CB, Cap AP, Spinella PC. Fresh whole blood transfusion for a combat casualty in austere combat environment. J.Spec.Oper.Med 2014;14(1):9-12.

27. Hakre S, Peel SA, O’Connell RJ, Sanders-Buell EE, Jagodzinski LL, Eggleston JC, Myles O, Waterman PE, McBride RH, Eader SA, et al. Transfusion-transmissible viral infections among US military recipients of whole blood and platelets during Operation Enduring Freedom and Operation Iraqi Freedom. Transfusion 2011;51(3):473-85.

28. Chan CM, Shorr AF, Perkins JG. Factors associated with acute lung injury in combat casualties receiving massive blood transfusions: a retrospective analysis. J.Crit Care 2012;27(4):419-14.

29. Hakre S, Manak MM, Murray CK, Davis KW, Bose M, Harding AJ, Maas PR, Jagodzinski LL, Kim JH, Michael NL, et al. Transfusion-transmitted human T-lymphotropic virus Type I infection in a United States military emergency whole blood transfusion recipient in Afghanistan, 2010. Transfusion 2013;53(10):2176-82.


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31. Schrager JJ, Branson RD, Johannigman JA. Lessons from the tip of the spear: medical advancements from Iraq and Afghanistan. Respir.Care 2012;57(8):1305-13.

32. Jenkins DH, Rappold JF, Badloe JF, Berseus O, Blackbourne L, Brohi KH, Butler FK, Cap AP, Cohen MJ, Davenport R, et al. Trauma hemostasis and oxygenation research position paper on remote damage control resuscitation: definitions, current practice, and knowledge gaps. Shock 2014;41 Suppl 1:3-12.

33. Berseus O, Boman K, Nessen SC, Westerberg LA. Risks of hemolysis due to anti-A and anti-B caused by the transfusion of blood or blood components containing ABO-incompatible plasma. Transfusion 2013;53 Suppl 1:114S-23S.

34. Strandenes G, Berseus O, Cap AP, Hervig T, Reade M, Prat N, Sailliol A, Gonzales R, Simon CD, Ness P, et al. Low titer group O whole blood in emergency situations. Shock 2014;41 Suppl 1:70-5.

35. Snyder EL, Whitley P, Kingsbury T, Miripol J, Tormey CA. In vitro and in vivo evaluation of a whole blood platelet-sparing leukoreduction filtration system. Transfusion 2010;50(10):2145-51.

36. AABB, Standards for Blood Banks and Transfusion Services. 30th edition. 2016.37. Spinella PC, Pidcoke HF, Strandenes G, Hervig T, Fisher A, Jenkins D, Yazer M,

Stubbs J, Murdock A, Sailliol A, et al. Whole blood for hemostatic resuscitation of major bleeding. Transfusion 2016;56 Suppl 2:S190-S202.

38. Polge C, Smith AU, Parkes AS. Revival of spermatozoa after vitrification and dehydration at low temperatures. Nature 1949;164(4172):666.


40. Pert JH, Schork PK, Moore R. Low-temperature preservation of human erythrocytes: biochemical and clinical aspects.Bibl.Haematol. 1964;19:47-53.



43. Valeri CR, Brodine CE. Current methods for processing frozen red cells. Cryobiology 1968;5(2):129-35.




47. Mazur P. Limits to life at low temperatures and at reduced water contents and water activities. Orig.Life 1980;10(2):137-59.

48. Mazur P. Stopping biological time. The freezing of living cells. Ann.N.Y.Acad.Sci. 1988;541:514-31.

49. Valeri CR, Ragno G, Pivacek LE, Cassidy GP, Srey R, Hansson-Wicher M, Leavy ME. An experiment with glycerol-frozen red blood cells stored at -80 degrees C for up to 37 years. Vox Sang. 2000;79(3):168-74.

50. UK National Blood Services , Guidelines for the Blood Transfusion Services in the UK, Change Notification UK National Blood Services No. 32 - 2016, 15 Aug 2016. 8th edition. 2013.

51. Council of Erope, Guide to the Preparation, Use and Quality Assurance of Blood Components. 18th edition. 2015. Strasbourg

52. Clinical and Practical Aspects of the Use of Frozen Blood. AABB; 1977.pp 23-3653. Valeri CR, Ragno G, Pivacek LE, Srey R, Hess JR, Lippert LE, Mettille F, Fahie

R, O’Neill EM, Szymanski IO. A multicenter study of in vitro and in vivo values in human RBCs frozen with 40-percent (wt/vol) glycerol and stored after deglycerolization for 15 days at 4 degrees C in AS-3: assessment of RBC processing in the ACP 215. Transfusion 2001;41(7):933-9.

54. Crowley JP, Wade PH, Wish C, Valeri CR. The purification of red cells for transfusion by freeze-preservation and washing. V. Red cell recovery and residual leukocytes after freeze-preservation with high concentrations of glycerol and washing in various systems. Transfusion 1977;17(1):1-7.


56. Hess JR, Hill HR, Oliver CK, Lippert LE, Greenwalt TJ. The effect of two additive solutions on the postthaw storage of RBCs. Transfusion 2001;41(7):923-7.

57. Moore GL, Ledford ME, Mathewson PJ, Hankins DJ, Shah SB. Post-thaw storage at 4 degrees C of previously frozen red cells with retention of 2,3-DPG. Vox Sang. 1987;53(1):15-8.


59. Neuhaus SJ, Wishaw K, Lelkens CCM. Australian experience with frozen blood products on military operations. Med.J.Aust. 2010;192(4):203-5.

60. Lelkens CCM, de Korte D, Lagerberg JWM. Prolonged postthaw shelf life of red cells frozen without prefreeze removal of excess glycerol. Vox Sang. 2015;108(3):219-25.


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63. Leitner GC, Dettke M, List J, Worel N, Weigel G, Fischer MB. Red blood units collected from bone marrow harvests after mononuclear cell selection qualify for autologous use. Vox Sang. 2010;98(3 Pt 1):e284-e289.


65. Valeri CR. Use of rejuvenation solutions in blood preservation. Crit Rev.Clin.Lab Sci. 1982;17(4):299-374.

66. Usry RT, Moore GL, Manalo FW. Morphology of stored, rejuvenated human erythrocytes. Vox Sang. 1975;28(3):176-83.

67. Raat NJH, Hilarius PM, Johannes T, de Korte D, Ince C, Verhoeven AJ. Rejuvenation of stored human red blood cells reverses the renal microvascular oxygenation deficit in an isovolemic transfusion model in rats. Transfusion 2009;49(3):427-4.

68. Gelderman MP, Vostal JG. Rejuvenation improves roller pump-induced physical stress resistance of fresh and stored red blood cells. Transfusion 2011;51(5):1096-104.

69. Koshkaryev A, Zelig O, Manny N, Yedgar S, Barshtein G. Rejuvenation treatment of stored red blood cells reverses storage-induced adhesion to vascular endothelial cells. Transfusion 2009;49(10):2136-43.

70. Barshtein G, Gural A, Manny N, Zelig O, Yedgar S, Arbell D. Storage-induced damage to red blood cell mechanical properties can be only partially reversed by rejuvenation. Transfus.Med Hemother. 2014;41(3):197-204.


72. FDA, Circular of Information for the Use of Human Blood and Blood Components, revised November 2013.

73. Hess JR, Lelkens CCM, Holcomb JB, Scalea TM. Advances in military, field, and austere transfusion medicine in the last decade. Transfus.Apher.Sci. 2013;49(3):380-6.

74. Fabricant L, Kiraly L, Wiles C, Differding J, Underwood S, Deloughery T, Schreiber M. Cryopreserved deglycerolized blood is safe and achieves superior tissue oxygenation compared with refrigerated red blood cells: a prospective randomized pilot study. J.Trauma Acute.Care Surg. 2013;74(2):371-6.

75. Holley A, Marks DC, Johnson L, Reade MC, Badloe JF, Noorman F. Frozen blood products: clinically effective and potentially ideal for remote Australia. Anaesth.Intensive Care 2013;41(1):10-9.

76. Hampton DA, Wiles C, Fabricant LJ, Kiraly L, Differding J, Underwood S, Le D, Watters J, Schreiber MA. Cryopreserved red blood cells are superior to standard liquid red blood cells. J.Trauma Acute.Care Surg. 2014;77(1):20-7.

77. Schreiber MA, McCully BH, Holcomb JB, Robinson BR, Minei JP, Stewart R, Kiraly L, Gordon NT, Martin DT, Rick EA, et al. Transfusion of cryopreserved packed red blood cells is safe and effective after trauma: a prospective randomized trial. Ann.Surg. 2015;262(3):426-33.

78. Dumont LJ, AuBuchon JP. Evaluation of proposed FDA criteria for the evaluation of radiolabeled red cell recovery trials. Transfusion 2008;48(6):1053-60.

79. Hess JR. Red cell freezing and its impact on the supply chain. Transfus.Med. 2004;14(1):1-8.

80. Noorman F, van Dongen TTCF, Plat MC, Badloe JF, Hess JR, Hoencamp R. Transfusion: -80 degrees C Frozen Blood Products Are Safe and Effective in Military Casualty Care. PLoS.One. 2016;11(12):e0168401.

Chapter

Summary

9

Chapter 9 Summary

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This thesis describes research into the possibilities to improve blood transfusion under battlefield conditions. Since the US Civil War (1861-1865), we have moved from person-to-person transfusion of whole blood, without any knowledge of blood groups, to highly sophisticated techniques, enabling preparation and storage of fully tested, ABO-Rhesus typed blood components for future use.

Driven by new tasks and experiences from mainly Bosnia, Iraq and Afghanistan, the Netherlands Military Blood Bank turned from a blood transfusion service into a top-notch, professional blood banking institute, ready to support military deployments worldwide. Nonetheless, despite technical developments over the past decades, like improved body armor, exsanguination still remains the major cause of preventable death on the battlefield. Hemorrhaging battlefield casualties call for a combination of quick, short, surgical intervention and early blood transfusion, the twofold mainstay of the so-called damage control resuscitation. This means that safe and effective blood should be made available at all times in the right amounts to surgical teams, as close to the point of injury as possible. Unpredictable characteristics of a battle zone leave little room for relying on ad-hoc donations from on-site (potentially unsafe) donors. Extending the shelf lives of red cells (and platelets) would enable a more efficient use and better availability of these scarce, invaluable commodities. To date, only freezing techniques can achieve considerably prolonged storage times, while retaining the desired properties of these essential blood components. After the first use of frozen red cells during the Vietnam War, the Netherlands military blood bank has, since the turn of the century, developed a full array of frozen blood components, needed to effectively treat massively bleeding patients. Together with a new transfusion protocol, survival rates now match those of civilian trauma centers. Nonetheless, some adjustments could improve the practical applicability of frozen red cells, particularly on the battlefield.

Chapter 1 reviews the developments in blood transfusion in the major military conflicts since the US Civil War. The rediscovery of the potential and application of citrate during the First World War brought about the possibility to store blood close to the point of care for 26 days. With the exception of a short-lived experiment with cadaver blood in the 1930s, the next major development was the fractionation of whole blood in 1940, yielding albumin to treat burn victims in particular. After initial enthusiasm, attention shifted back again to whole blood and that didn’t change until the 1960s and 1970s, when component therapy became the standard.

After the accidental discovery of glycerol as a useful cryoprotectant, the US started the development of a frozen red cell program in the late 1950s, leading to deployment of frozen blood banks in the late 1960s during the Vietnam War. Several drawbacks, however, led to abolition of these blood banks.

During the past two decades, experiences obtained from the various military conflicts have learnt that the vast majority of standard red cell concentrates had to be discarded due to outdating.

The only storage method, currently available, to minimize wastage due to expiration, is the application of cryopreservation. The Netherlands’ extensive, favorable experiences with frozen blood products during missions in Southwest Asia was the basis for further research in attempts to improve the practicability of the use of frozen red cells in battlefield conditions.

Chapter 2 compares the effects on postthaw preservation of the two main methods of cryopreserving red cells, the high- (40%) glycerol method (HGM) and the low- (19%) glycerol method (LGM). Glycerol treatment by itself induced hemolysis during processing, which was more pronounced in HGM cells. The freeze-thaw-wash process in a fully closed system (ACP215, Haemonetics) is indeed a prerequisite for prolonged postthaw storage, but decreased the stability of RBCs, particularly in LGM cells during storage after thawing. The results showed lower hemolysis in HGM red cells than in LGM red cells. It was concluded that the closed washing system is able to process both high- and low-glycerol-treated RBCs. Stability after washing during cold storage in SAGM, as measured by hemolysis, is better for HGM cells as compared to LGM cells.

Based on these methods and results, in chapter 3 the Netherlands’ experiences with (universal) - 80⁰C frozen blood products in military trauma settings are discussed, with special attention to quality control. All thawed (washed) blood products proved to be in compliance with international regulations and guidelines. The most important conclusions were that the -80°C frozen stock of HGM red cells, plasma and platelets, readily available after thaw (and wash), enables a reduction of shipments, as well as the abolition of the backup “walking blood bank”, without compromising the availability of safe blood products near the battlefield.

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Chapter 4 reports the experiences from an Australian perspective with the Netherlands’ frozen blood products to treat major trauma victims. Historically, the Australian Defence Force (ADF) has sourced all its blood supplies from the Australian Red Cross Blood Service. For the first time, the ADF relied on another nation’s blood supply system. In 2008, the ADF embedded a surgical and intensive care team into the Netherlands-led forward health facility at Tarin Kowt in Uruzgan, Afghanistan. The course of events in two hemorrhaging casualties from admission to discharge/transfer is reported. Despite higher costs, the integrated fresh–frozen blood banking facility provided flexible and efficient use of blood products in a military setting.

Chapter 5 addresses the influence of supernatant prefreeze glycerol reduction on the stability of thawed, deglycerolized RBCs during subsequent cold storage. Currently, glycerolization (ACP215, Haemonetics) is followed by supernatant glycerol reduction before freezing to decrease the volume of the frozen red cells. The aim of this study was to investigate the influence of skipping supernatant glycerol reduction before freezing on the stability of thawed, deglycerolized RBCs during subsequent cold storage. After storage at -80°C, the units were thawed, deglycerolized (ACP215, Haemonetics) and resuspended in SAGM or AS-3. During cold storage (2-6°C), the red cells were analyzed for their stability and in vitro quality. The freeze-thaw-wash recovery proved to be comparable for both volume reduced and non-reduced units. During postthaw storage, non-glycerol reduced units showed significantly less potassium leakage and hemolysis and higher ATP levels. As compared to SAGM, AS-3 strongly reduced hemolysis during postthaw storage of non-glycerol reduced units: hemolysis remained below 0.8% for up to 28 days of storage. Omitting the glycerol supernatant reduction before freezing not only simplifies the cryopreservation procedure, but also increases the stability, and therefore the outdating period of thawed RBCs. This increases the practical applicability of cryopreserved RBCs in both civil (rare blood) and military blood transfusion practice.

Chapter 6 describes the effect of prefreeze rejuvenation on postthaw storage of red cells in two additive solutions, AS-3 and SAGM. We postulated that using red cells with increased ATP and 2,3-DPG content, prior to freezing could extend their postthaw shelf life, while still meeting hemolysis (0.8%) and total adenylate (> 82% of original values) requirements. To reach the necessary higher

levels of total adenylate and 2,3-DPG, RBC units (in SAGM) were incubated with Rejuvesol and, for comparison, RBC units were stored in PAGGGM, an additive solution designed to increase metabolic status. After glycerolization without prefreeze volume reduction (result from chapter 5) red cell units were frozen and stored for at least 14 days at -80°C.After deglycerolization, cells were resuspended in SAGM or AS-3. Based on a maximum allowable hemolysis of 0.8% and a total adenylate content of > 82% of the original value, thawed, prefreeze Rejuvesol or PAGGGM red cell units can be stored for 35 days at 2-6ºC in AS-3. This means that thawed, previously frozen red cells have a shelf life, comparable to the one of standard liquid red cells, with better metabolic characteristics (especially 2,3-DPG levels) during the first weeks of storage at 2-6⁰ C. In addition to the results obtained in chapter 5, these findings further improve the practical applicability of frozen red cells in both civil (rare blood groups) and military blood transfusion practice.

Chapter 7 reviews the advances in military, field, and austere transfusion medicine in the last decade, based on the experiences of twenty years of war in Southwest Asia. The wars in the Gulf, Iraq and Afghanistan have demonstrated the essential role of primary resuscitation with blood products in the care of critically injured soldiers. This idea has been widely adopted and is being critically tested in civilian trauma centers. The need for red cells, plasma and platelets to be immediately available in remote locations creates a logistic burden that will best be eased by innovative new blood products such as longer-stored liquid RBCs, freeze-dried plasma, small-volume frozen platelets, and coagulation factor concentrates such as fibrinogen concentrates and prothrombin complex concentrates. Such products have long shelf lives, low logistic burdens of weight, fragility, or needs for processing prior to use. Developing and fielding a full family of such products will improve field medical care and make products available in the evacuation chain. It also will allow treatment in other austere environments such as the hundreds of small hospitals in the US which serve as Levels 3 and 4 trauma centers but do not currently have thawed plasma or platelets available. Such small trauma centers currently care for half of all the trauma patients in the country. Proving the new generation of blood products work, will help assure their widest availability in emergencies. Currently, the Dutch are the only country in the world with extensive experience in fielding and using the full array of frozen blood products. They used them extensively with excellent results in

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Iraq and Afghanistan and so did the Australians, who depended on the Dutch blood supply in Afghanistan. Finally, the background and reasons for diverting to a different preservation technique for military purposes and improvements therein are discussed in Chapter 8.

Chapter

Samenvatting

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Dit proefschrift beschrijft onderzoek naar mogelijkheden om de praktische toepasbaarheid van diepgevroren rode bloedcellen onder met name oorlogsomstandigheden te vergroten.

Sinds de Amerikaanse Burgeroorlog (1861-1865) heeft bloedtransfusie zich de afgelopen 150 jaar ontwikkeld van een directe overbrenging van volbloed van donor naar ontvanger, zonder enige kennis van bloedgroepen, naar een hoogtechnologische aangelegenheid, waarbij volledig geteste bloedcomponenten, gesorteerd naar bloedgroep, worden bereid en opgeslagen onder optimale condities voor uitgesteld gebruik.

Na de val van de Berlijnse Muur in 1989 kreeg de Nederlandse krijgsmacht als nieuwe taak, vastgelegd in de Grondwet, het wereldwijd helpen bevorderen van de internationale rechtsorde. In de praktijk houdt dit deelname aan (militaire) missies, ver buiten Nederland, in, waarbij, naast doden, rekening dient te worden gehouden met grotere aantallen ernstig gewonden met veel bloedverlies. Ondanks technologische ontwikkelingen, zoals scherf- en kogelwerende lichaamsbescherming, is verbloeding nog steeds de belangrijkste, vermijdbare doodsoorzaak bij militairen op het slagveld. Hierbij is onvoorspelbaar wanneer op welke plaats hoeveel bloedproducten nodig zijn. Dit proefschrift richt zich voornamelijk op de benodigde rode bloedcellen. Deze cellen kennen, afhankelijk van het gebruikte bewaarmedium en/of toegepaste regelgeving, een bewaartijd van 35 tot 42 dagen. Dit betekent dat, onafhankelijk van verbruik, in ieder geval één keer per maand een voorraad rode bloedcellen moet worden ververst. Zelfs als de operationele omstandigheden zodanig zijn dat herbevoorrading onbelemmerd kan plaatsvinden, zijn er nog altijd de factoren tijd en afstand die, samen met de minimaal noodzakelijke frequentie van herbevoorrading, het gebruik van standaard rode bloedcellen bemoeilijken. Het onvoorspelbare verbruik leidt enerzijds gemakkelijk tot verspilling, anderzijds moet worden gevreesd voor tekorten op bepaalde tijdstippen. Naast het feit dat dit onnodig veel financiële middelen vereist, is een tekort operationeel ongewenst en een verspilling ethisch moeilijk te verantwoorden naar de donoren, die uit ideële overwegingen om niet hun bloed ter beschikking stellen.

Aangezien veel missies een looptijd kennen van meerdere jaren, betekent dit dat operationele en logistieke, maar ook financiële en ethische redenen het gebruik van standaard rode bloedcellen als onderdeel van een noodzakelijke transfusiebehandeling ongewenst maken. De oplossing ligt in het gebruik van veel langer houdbare producten, die gedurende hun opslag in een inzetgebied

langdurig hun effectiviteit behouden en ter plaatse, indien nodig, gemakkelijk in korte tijd geschikt gemaakt kunnen worden voor toediening. Dit betekent voor de hedendaagse praktijk het gebruik van diepvriesmethoden. De aanzet hiervoor werd gegeven in 1949, toen na een toevallige ontdekking bleek dat glycerol een bruikbaar middel was om rode bloedcellen in te vriezen, zonder deze al te zwaar te beschadigen. Een belangrijk nadeel is echter dat het gebruikte glycerol weer moet worden verwijderd tot een zeer lage concentratie vóór toediening, omdat anders de toegediende rode bloedcellen opzwellen en kapotgaan, wat voor de patiënt uiterst nadelige consequenties heeft. Dit vergt niet alleen kostbare tijd, maar ook geld, vanwege de benodigde apparatuur. Deze factoren staan, samen met de beperkte houdbaarheid van 14 dagen na ontdooien en wassen, een grotere flexibiliteit en slagvaardigheid bij het gebruik van diepgevroren rode bloedcellen, zeker onder primitieve omstandigheden als in een militair operatiegebied, in de weg.

Het eerste militaire gebruik van diepgevroren rode bloedcellen in glycerol dateert uit de tweede helft van de jaren ‘60 in Vietnam. De hierbij gebruikte techniek stond voor de Nederlandse Militaire Bloedbank rond de overgang naar de 21e eeuw aan de basis van de ontwikkeling van een compleet assortiment diepgevroren bloedproducten om massaal bloedende patiënten succesvol te kunnen behandelen. Samen met een nieuw ontwikkeld transfusieprotocol heeft dit uiteindelijk geleid tot overlevingspercentages die vergelijkbaar zijn met die van civiele traumacentra. Niettemin kunnen enkele aanpassingen in de gebruikte techniek de praktische toepasbaarheid van diepgevroren rode bloedcellen verder verbeteren, m.n. onder gevechtsomstandigheden. De onderzoeken die hiervoor zijn uitgevoerd, worden in dit proefschrift beschreven.

In hoofdstuk 1 komen als eerste de ontwikkelingen in bloedtransfusie aan bod in enkele grote militaire conflicten sinds de Amerikaanse Burgeroorlog. Uiteindelijk wordt in de jaren 1960-70 bloedcomponententherapie de standaard, gebaseerd op de overwegingen dat met hetzelfde bloed zoveel mogelijk mensen geholpen moeten kunnen worden en de bewaaromstandigheden voor elke component geoptimaliseerd kunnen worden. In de verschillende militaire conflicten gedurende de afgelopen twee decennia, bleek dat het overgrote deel van de voorraden rode bloedcellen ongebruikt moest worden weggegooid. Nederland heeft in die periode uitgebreid ervaring opgedaan met het gebruik van diepgevroren rode cellen in met name Irak en Afghanistan, wat de basis vormde voor nader onderzoek. Voorop stond hierbij het verbeteren van de


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praktische toepasbaarheid van bevroren rode bloedcellen in militair operationele omstandigheden.

Zo beschrijft hoofdstuk 2 de twee belangrijkste methoden van diepvriezen van rode bloedcellen, met als conclusie de bevestiging dat de voor militair gebruik meest toegankelijke methode ook de meest stabiele rode cellen oplevert na ontdooien en wassen.

Uitgaande van deze methode laat hoofdstuk 3 zien dat in de militaire praktijk niet alleen alle ontdooide, gewassen eenheden rode bloedcellen aan de internationaal geldende voorschriften voldoen, maar ook dat het gebruik ervan de aantallen benodigde herbevoorradingen kan reduceren. Daarnaast maakt dit systeem een “walking blood bank”, bestaande uit lokaal personeel dat vrijwillig doneert, overbodig.

Aan de hand van twee casussen beschrijft hoofdstuk 4 vervolgens vanuit Australisch perspectief het verloop van de behandeling met uit Nederland afkomstige diepgevroren bloedproducten, waaronder rode bloedcellen. De conclusie luidt dat, ondanks hogere investeringen, een voorraad van ontdooide en gewassen rode bloedcellen, samen met een diepgevroren hoeveelheid, een flexibel en efficiënt gebruik van bloedproducten in een militaire omgeving mogelijk maakt.

Een belangrijke beperking van het gebruik van diepgevroren bloedproducten is de beperkte houdbaarheid na ontdooien. Er is daarom onderzocht of het, door aanpassing van de vriesmethode, mogelijk is deze te verlengen.

Een eerste aanpassing is het gebruik van een wasvloeistof met een neutrale pH, die al eerder werd beschreven door onderzoekers van Sanquin, waarmee ook andere aanpassingen, zoals beschreven in dit proefschrift, werden uitgewerkt. In hoofdstuk 5 wordt deze modificatie als uitgangspunt gebruikt en gecombineerd met een tweede aanpassing, waarbij het oorspronkelijke volume van de eenheid in glycerol niet verkleind wordt vóór invriezen. Dit onderzoek laat zien dat de combinatie van deze wijzigingen in de procedure minder beschadiging van de rode cellen tot gevolg heeft, waardoor ze, na ontdooien, in plaats van de huidige twee weken, 28 dagen bruikbaar blijven voor transfusie. Indien de rode cellen vóór invriezen bovendien worden behandeld met stoffen die kunstmatig hun energievoorraad verhogen tot boven de normale niveaus, wordt hun houdbaarheid na ontdooien nog verder verlengd tot 35 dagen, zo laat hoofdstuk 6 zien. Daarmee is een houdbaarheid bereikt die gelijk is aan die van standaard rode bloedcellen, zoals die normaliter in de burgermaatschappij

worden gebruikt. Hoofdstuk 7 blikt terug op de ontwikkelingen op het gebied van bloedtransfusie onder primitieve omstandigheden, m.n. onder militaire condities, gedurende de afgelopen tien jaar. Vertrekpunt hierbij is het geheel aan ervaringen, opgedaan op dit gebied gedurende de afgelopen twintig jaar van oorlog, m.n. in Irak en Afghanistan. De rol van bloedproducten in de primaire behandeling van massaal bloedende slachtoffers is essentieel gebleken en dat heeft mede geleid tot aanpassing van procedures in civiele traumacentra. De logistiek blijft echter een groot probleem, m.n. civiel op afgelegen locaties en onder militaire omstandigheden. Om dit op te lossen bestaat er een algemene, brede behoefte aan nieuwe producten met een langdurige houdbaarheid onder eenvoudige bewaarcondities zonder noodzakelijke, ingewikkelde bewerkingen vóór transfusie. Tot op heden zijn diepgevroren bloedproducten wat betreft lange houdbaarheid en effectiviteit het best beschikbare alternatief gebleken voor Nederland. Ten slotte belicht hoofdstuk 8 in een algemene discussie eerst de achtergrond en de redenen om te kiezen voor het diepvriezen van rode cellen ten behoeve van militair operationeel gebruik. Aansluitend volgt dan de bespreking van de onderzoeksresultaten uit dit proefschrift, die aanzienlijke verbeteringen hebben laten zien in de houdbaarheid en kwaliteit van ontdooide rode bloedcellen.

Dankwoord / Acknowledgments

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Dankwoord / Acknowledgments

De reis die, vanaf een eerste kennismaking met bloedtransfusie in de krijgsmacht, via de gedachte over een promotie, naar de totstandkoming van een proefschrift leidde, heeft mij in aanraking gebracht met veel bijzondere mensen. Allemaal hebben zij, op enigerlei wijze, met hun kennis, steun en ervaring mij richting eindstreep geholpen. Zonder de pretentie volledig te zijn, is hier een woord van dank daarom zeer zeker op zijn plaats.

Prof. dr. A.J. Verhoeven, mijn promotor: Beste Arthur, we hebben, wat mij betreft, een ongebruikelijk promotiepad bewandeld. Ik wilde graag de mogelijkheid onderzoeken om een aantal al gepubliceerde artikelen te gebruiken als basis voor een promovabel geheel, waarbij ik nog iemand zocht die als promotor wilde optreden. Al heel snel kwam ik vervolgens bij jou uit, omdat ik vond dat je, ook gezien je achtergrond en verleden bij Sanquin, helemaal paste bij het onderwerp van mijn keuze. Bovendien had ik goede herinneringen aan een eerste gezamenlijke publicatie uit de tijd dat ik net begonnen was bij de Militaire Bloedbank. Ik prijs mij gelukkig dat jij, mede op advies van beide copromotoren, je bereid hebt verklaard de taak van promotor op je te nemen. Dit helpt trouwens ook het gevoelige punt te verzachten, dat een geboren Leienaar en alumnus van ’s lands oudste universiteit, uiteindelijk promoveert in Amsterdam.

Dr. J.W.M. Lagerberg, copromotor: Beste Johan, de afgelopen jaren hebben we intensief contact gehad over het opzetten van nieuw onderzoek rond diepgevroren rode bloedcellen en de uitwerking van de bijbehorende resultaten in artikelen. Dank dat je, ondanks alle dagelijkse, drukke, andere bezigheden en beslommeringen, toch nog altijd ergens tijd vond om mijn ideeën aan de wetenschappelijke mores te toetsen, zowel via de e-mail, de telefoon als in de besprekingen in Amsterdam. Het was uiterst plezierig met je samen te werken!

Dr. D. de Korte, copromotor: Beste Dirk, ik herinner mij nog dat ik eind 2013 besloot om jou te benaderen met de vraag of ik onder de vleugels van Sanquin zou kunnen promoveren. Na je goedkeuring toonde je je sindsdien een altijd zeer kritisch beoordelaar van de concepten, die ik ter bespreking aanbood. Er VN-missie Cambodja, Veldhospitaal te Phum Nimith, 1993

Appendices Dankwoord / Acknowledgments

166 167

was altijd iets dat toch nog beter kon, als ik mij weer eens had blind gestaard op mijn werkstuk. Mijn dank voor je niet-aflatende kritische blik, die mede tot dit proefschrift heeft geleid. Ik heb de samenwerking zeer gewaardeerd!

Dr. F. Noorman: Beste Femke, je vaardigheden als onderzoeker en kwaliteits-functionaris kwamen bij de Militaire Bloedbank meer dan tot hun recht. Ik heb je leren kennen en waarderen als een enthousiaste, onconventionele denker, die de (militaire) mogelijkheden van diepgevroren bloedproducten internationaal naar de top heeft gebracht. Het bewijs daarvan hebben we op menig congres als poster of lezing naar voren gebracht. Ik denk met plezier terug aan de gesprekken in de deurpost tussen onze beide kamers in Leiden, waarbij we brainstormden over de verbetering van vooral de was- en resuspensieprocedure van ontdooide rode bloedcellen. Jouw enthousiasme en bijdragen hebben mede aan de wieg gestaan van dit proefschrift, waarvoor mijn dank.

Prof. dr. A. Brand: Beste Anneke, jij bent mijn leermeester in de transfusie-geneeskunde geweest. In mijn beleving was wat jij niet wist ook de moeite van het weten niet waard. Naast het klinische werk werd “Mollison” gedurende de opleiding van kaft tot kaft (een hoofdstuk per week) doorgenomen, waardoor ik goed beslagen ten ijs kon komen in de V.S. voor het allerlaatste jaar van mijn opleiding. Mijn oprechte dank voor jouw bijdrage aan de ontwikkeling van mijn kennis en ervaring op het gebied van bloed en bloedproducten, ook en vooral bij de behandeling van patiënten.

Prof. dr. C. Th. Smit Sibinga: Beste Cees, jarenlang vertoefde ik op jouw uitnodiging enkele dagen in het hoge noorden, om daar het door jouw (toenmalige) Rode Kruis Bloedbank Groningen-Drenthe georganiseerde wetenschappelijke symposium bij te wonen. Deze symposia, samen met jouw persoonlijke inzet, hebben in belangrijke mate bijgedragen aan mijn kennisontwikkeling van bloedproducten en bijbehorende technologieën. Ik heb je bijzondere betrokkenheid bij mijn militaire werk en het aandeel van bloedtransfusie daarin altijd bijzonder gewaardeerd. Dank daarvoor!

C.R. Valeri, CAPT (ret.), MC, USN: Dear Bob, in 1987 I sought your advice on how to use frozen blood products in the military. My first visit in 1988 to your Naval Blood Research Laboratory in Boston left a lasting impression for the

rest of my military career. I kept pursuing your ideas to get them implemented into our military blood supply system and we have kept in touch ever since. In 2001, your laboratory tested our frozen platelets and the results proved to fill in a void in the operational medical treatment system during deployments. Over the past decade, our frozen blood products have helped saving numerous lives in particularly Iraq and Afghanistan, without serious adverse events or unnecessary exposure to transfusion transmissible diseases. You were the shoulders we stood on. Thank you!

Prof. J.R. Hess, MD, MPH, FACP, FAAAS: Dear John, we go back a long way. When we first met, you were still a colonel in the US Army. During our conversations at AABB conferences, my SBB training at Walter Reed Army Medical Center and fellowship at R Adams Cowley Shock Trauma Center in Baltimore, you were the tutor who filled the gaps in my knowledge with regard to treating massively bleeding patients. Thank you and thank you also for accepting a seat on the Doctorate Committee.

Overige leden van de promotiecommissie: Ik waardeer niet alleen uw belang-stelling voor dit proefschrift, maar ook het feit dat u zitting heeft willen nemen in de promotiecommissie.

Niet te vergeten, de analisten Erik Gouwerok, Mya Go en Richard Vlaar. Zonder jullie bijdrage zou er letterlijk geen resultaat op papier zijn verschenen. Dank voor het mij wegwijs maken in de principes van de verschillende analyses en de uitvoering van de diverse bepalingen. Mede dankzij jullie kan ik terugzien op enkele geaccepteerde posters en artikelen. Mijn dank en waardering voor jullie bijdrage is groot!

Jacqueline, mijn onvolprezen echtgenote, die mij stimuleerde om na mijn functioneel leeftijdsontslag de sluimerende promotieplannen versneld weer op te pakken. Al jaren vraag je mij om nu eindelijk eens de garage op te ruimen, zodat deze gebruikt kan worden waarvoor hij bedoeld is en mijn studeerkamer te veranderen in een gezellig verblijf zonder stapels papier, die op tafels, in stoelen en op de grond verspreid liggen. Je kunt gerust zijn: ik kan en ga nu eindelijk mijn belofte inlossen.

Curriculum vitae auctoris

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Appendices

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Curriculum vitae auctoris

De schrijver van dit proefschrift werd op 16 september 1954 geboren in Leiden. In aansluiting op het eindexamen Gymnasium B in 1972 aan het Bernardinuscollege te Heerlen begon hij zijn studie geneeskunde aan de Rijksuniversiteit te Leiden. Na zijn afstuderen in 1979 vervulde hij zijn militaire dienstplicht bij de Koninklijke Marine bij het Marine Keurings- en Selectiecentrum te Hilversum en aan boord van het standaardfregat Hr.Ms. Banckert. Een contract als beroepsofficier voerde hem vervolgens tot januari 2010 gedurende bijna dertig jaar langs een breed scala aan functies. Zo was hij bataljonsarts bij de Eerste Amfibische Gevechtsgroep van het Korps Mariniers te Doorn, Hoofd Geneeskundige Dienst van de Zeemacht in het Caraïbisch gebied op Curaçao en stafarts van het Korps Mariniers te Rotterdam. Ook werd hij uitgezonden naar Cambodja (1992-1993), voormalig Joegoslavië (1997-1998) en Afghanistan (2007). Tussendoor behaalde hij een propedeuse in de rechten aan de Rijksuniversiteit te Utrecht (1990), rondde hij de opleiding af tot specialist Algemene Gezondheidszorg te Leiden (1993) en was hij, vanuit het Nederlandse Ministerie van Defensie, projectleider rehabilitatie van het militair hospitaal te Paramaribo (1996-1997). Een driejarige opleiding (1998-2001) tot transfusiearts bij achtereenvolgens de Bloedbank Leiden-Haaglanden, het Leids Universitair Medisch Centrum en het Walter Reed Army Medical Center te Washington D.C. vormde de voorbereiding op zijn laatste functie bij Defensie, die van commandant van de Militaire Bloedbank. Direct na de afronding van deze opleiding werd hij in 2001 ingeschreven als Specialist in Blood Banking (SBB) van de American Society for Clinical Pathology (ASCP). Onder leiding van prof. John R. Hess volgde hij in 2004 een aanvullende opleiding, gericht op het gebruik van bloedproducten bij de behandeling van (zeer) groot trauma, in het R Adams Cowley Shock Trauma Center in Baltimore, MD, V.S. In 2007 volgde zijn benoeming tot Adjunct Assistant Professor aan de George Washington University te Washington D.C. en certificering als assessor van de American Association of Blood Banks (AABB). Na zijn functioneel leeftijdsontslag in de rang van kapitein ter zee-arts in december 2009 heeft hij, onder leiding van prof. dr. C. Th. Smit Sibinga, in 2011 en 2012 deelgenomen aan verkennende missies naar Kazachstan en Kirgizië, gericht op verbetering van de lokale bloedvoorziening in die landen. Hij is sinds 1987 getrouwd met Jacqueline en vader van Marie-Christine (1991) en Jean-Louis (1994).

Arts bij het Korps Mariniers (Happy van der Valk Bouman, Curaçao, 1986)

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Resume

Resume

The author of this thesis was born on September 16, 1954, in Leiden, the Netherlands. He graduated from Leiden University medical school in 1979 and joined the Royal Netherlands Navy as a lieutenant (junior grade) in 1980. He obtained his first-year degree in Dutch law in 1990 from the Utrecht University and completed a study in community medicine and public health in Leiden in 1993.In his capacity as the Staff Medical Officer of the Royal Netherlands Marine Corps (1990-1995) he commanded a Netherlands Armed Forces field hospital during the United Nations mission in Cambodia (1992-1993). He was also deployed to Former Yugoslavia (1997-1998) and Afghanistan (2007).In preparation for his last assignment in his military career, commanding officer of the Netherlands Military Blood Bank, he was trained in transfusion medicine and blood banking at the Leiden University Medical Center (1998-2000) and at Walter Reed Army Medical Center (2000-2001) in Washington D.C. Shortly after his graduation in 2001, he became board certified as a Specialist in Blood Banking (SBB) of the American Society for Clinical Pathology (ASCP). In 2004, as a fellow under the auspices of Prof. John R. Hess, he received additional training in the use of blood products in severe trauma at the R Adams Cowley Shock Trauma Center in Baltimore, MD, USA. He was appointed adjunct assistant professor of pathology at the George Washington University in Washington D.C. in 2007.From 2003 until his retirement from the Royal Netherlands Navy in the rank of Captain in December 2009, he chaired the NATO working group on blood and blood products, primarily aimed at mutual acceptance and interchangeability of blood products within the NATO membership states.Since 1987 he is married to his wife Jacqueline with two children, Marie-Christine (1991) and Jean-Louis (1994).

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