W.J.A. Dekker

Platelet activation during preparation of platelet concentrates: a comparison of the platelet-rich plasma and the buffy coat methods

The activation of platelets during the preparation of platelet concentrates (PCs) by two methods was compared. To eliminate interdonor differences, 2 units of whole blood were pooled and subsequently divided into two batches. From one batch, the platelets were harvested as pelleted platelets from platelet‐rich plasma (PRP) and from the other as nonpelleted platelets from the buffy coat (BC). The activation of platelets in these PCs was studied immediately after preparation and during storage for up to 9 days at 22°C with gentle agitation. The binding of monoclonal antibodies (MoAbs) against the GP IIb/IIIa complex and against activation‐dependent antigens (GMP 140 from the alpha granules and a 53‐kDa glycoprotein from the lysosomal granules) was measured. β‐thromboglobulin (β‐TG) release was also determined. Disc‐to‐sphere transformation was quantitated by measuring on an aggregometer the difference in light transmission during stirring at different rates and also by light microscopy. Immediately after preparation, platelets derived from PRP had a more spheric morphology (p < 0.01), had a higher β‐TG release (p < 0.01), bound more MoAbs against GP IIb/IIIa (p < 0.01), and expressed more GMP 140 and 53‐kDa glycoprotein (p < 0.01) than did BC‐ derived platelets. However, these differences had disappeared after 2 days of storage. It was concluded that, immediately after preparation, PRP‐derived platelets are more activated than BC‐derived platelets. This is most likely a result of the pelleting that follows the second high‐speed centrifugation of the PRP.

Preparation of Leukocyte-Poor Platelet Concentrates from Buffy Coats

Abstract. A special insert was developed for centrifuge cups in order to prepare leukocyte‐poor platelet concentrates from buffy coats by using quadruple citrate phosphate dextrose‐saline adenine glucose mannitol systems from different manufacturers. Each centrifuge cup could contain up to 4 sets of double bags allowing the preparation of 24 platelet concentrates per run. Optimal conditions for centrifugation of the buffy coats in the inserts were found to be 6 min at 380 g (2,150 g min). A platelet count of 69 ± 19 × 109 and a leukocyte contamination of 14± 10.5 × 106 per platelet concentrate was thereby obtained in a plasma volume of 63 ± 10.5 ml (mean ±SD). The method described allows large scale production of leukocyte‐poor platelet concentrates from buffy coats in a closed system.

Prevention of Yersinia enterocolitica growth in red-blood-cell concentrates

In response to concern about Yersinia enterocolitica contamination of blood products, we have studied the effects on Y enterocolitica growth of holding whole blood at 22 degrees C for 20 h and then removing leucocytes. Thirty pools of three bags of blood were inoculated with Y enterocolitica (2 x 10(1)-3 x 10(4) colony-forming units/ml). One bag in each pool was processed to red-blood-cell concentrate after 6 h at 4 degrees C (RBC); the other two were held at 22 degrees C for 20 h before processing to buffy-coat-depleted RBC (BCd-RBC). One of these bags was then depleted of leucocytes by filtration (Ld-RBC). All bags were stored at 4 degrees C for 5 weeks. RBC bags showed Y enterocolitica growth after the shortest storage times, followed by BCd-RBC then Ld-RBC (p less than 0.03-0.001). We recommend that whole blood should be held at 22 degrees C to make use of inherent bactericidal activity; leucocytes should then be removed.

Storage of leukocyte-poor red cell concentrates: filtration in a closed system using a sterile connection device

Abstract. Storage of leukocyte‐poor red cell concentrates (LP‐RCC) was investigated after filtration in a closed system that was assembled using a Sterile Connection Device (SCD). The LP‐RCC were stored for up to 6 weeks following filtration with either 0.9% saline solution (n = 14) or saline‐adenine‐glucose‐mannitol (SAG M) solution (n = 15) to prime and rinse the cellulose acetate filter. The results were compared with the data of nonfiltered buffy‐coat‐poor red cell concentrates (BC‐poor RCC) stored in SAG M solution (n = 10). All LP‐RCC contained less than 106 leukocytes whereas the nonfiltered BC‐poor RCC contained 6751286 × 106 leukocytes at day 1, decreasing to 83±49 × 106 at day 42. Although glucose consumption, lactic acid production and decrease in pH was similar from day 7 through 28 in both groups of LP‐RCC, a significantly steeper decline of ATP values as well as a higher hemolysis and LDH release was observed in the LP‐RCC filtered with saline. During storage of the nonfiltered BC‐poor RCC in SAG M, significantly higher glucose consumption (p<0.01), LDH release (p<0.001), rate of hemolysis (p<0.001) and a lower pH (p<0.001) were found, compared to the filtered units. It is postulated that the leukocytes present in the nonfiltered BC‐poor RCC were responsible for these differences. The ATP values in the SAG‐M‐filtered and nonfiltered BC‐poor RCC in SAG M were comparable. By comparing the ATP levels and values of the filtered RCC and the nonfiltered BC‐poor RCC we conclude that the LP‐RCC can be stored for 35 days if SAG M solution is used to prime and rinse the filter.

Preparation of leukodepleted platelet concentrates from pooled buffy coats: prestorage filtration with Autostop BC

Background and Objectives: Our requirements for leukocyte–depleted platelet concentrates (LD–PC) for an adult patient are: platelets >240×109, leukocytes <5×106, volume of 150–400 ml; and at the end of storage a pH between 6.8 and 7.4 and presence of the swirling effect. Our aim was to develop a standardized, semiautomated method for the production of LD–PC, by pooling of buffy coats (BC), and prestorage leukoreduction by filtration. Materials and Methods: Whole blood was collected in Top and Bottom systems, and separated automatically with the Compomat™ G3 equipment into a red cell concentrate, a plasma and a BC. Subsequently, a pool of 5 BC was made, and 200 g plasma from one of the donors was added. Then, after soft spin centrifugation, the platelet rich plasma was leukocyte depleted by filtration using the Autostop™BC filter, and stored in a 1,000 ml polyolefin platelet storage bag. Results: BC (n = 60) had a volume of 51±2 ml (mean ± SD) with a hematocrit of 0.44±0.03 l/l and contained 80±5% of the platelets and 74±12% of the leukocytes of the whole blood. Routinely prepared LD–PC (n = 15,037) contained a median of 341×109 platelets (range 49–599×109), with only 104/15,037 (0.7%) containing fewer than 240×109 platelets; the median volume was 263 ml (range 134–373 ml). In 118/917 (13%) LD–PC leukocytes were observed in the Nageotte hemocytometer, but only twice exceeding 1×106 leukocytes per unit, and none exceeding 5×106 (median <0.6×106; range <0.6–1.41×106). Storage experiments of the LD–PC (n = 12) revealed adequate oxygenation and maintenance of pH and swirling effect up to 9 days. Conclusions: This method warrants with 99% confidence that LD–PC contain more than 240×109 platelets; with 97.5% confidence that 100% of the LD–PC contain <5×106 leukocytes, and with 95% confidence that more than 99% of the LD–PC contain fewer than 1×106 leukocytes; these LD–PC can be stored satisfactorily for up to 9 days.

Comparison of five different filters for the removal of leukocytes from red cell concentrates.

The leukocyte depletion capacity and performance of 5 filters designed for filtration of red cell concentrates (RCC) were compared by counting leukocytes, measuring red cell volumes and by histological examination of the filters after use. To eliminate interdonor differences, 5 buffy‐coat‐poor RCC were pooled (in each of 10 experiments) and subsequently split up into the original bags. The RCC were passed over the Cellselect filter, a column filled with cellulose acetate, and over flat‐bed polyester filters: the Cellselect Optima, the Pall RC 50, the Leukostop and the Sepacell R‐500.

WBC‐reduced platelet concentrates from pooled buffy coats in additive solution: an evaluation of in vitro and in vivo measures

BACKGROUND: The use of a platelet additive solution (PAS‐II, Baxter) may have benefits over plasma for storage of platelets. It was the aim of this study to develop a method to produce WBC‐reduced platelet concentrates (PCs) in PAS‐II with >240 × 109 platelets and <1 × 106 WBCs per unit, which can be stored for 5 days at pH >6.8 and that will give sufficient platelet increments after transfusion: a 1‐hour CCI of >7.5 and a 20‐hour CCI of >2.5.

Comparison of a Conventional Quadruple-Bag System with a ‘Top-and-Bottom’ System for Blood Processing

‘Top‐and‐bottom’ bags have an outlet at the top and at the bottom of the collecting bag, allowing simultaneous expression of plasma and red cells, whereas the buffy coat remains in the collecting bag. The composition of blood components was investigated following manual separation of whole blood in a conventional 4‐bag system (A) or automated separation in a ‘top‐and‐bottom’ system (B). To overcome inter‐donor differences, two units of whole blood were pooled and redistributed into the original bags (A and B) prior to centrifugation. Leukocyte‐poor platelet concentrates (LPPC) were manufactured from both types of buffy coat (A and B). The volumes of plasma, red cell concentrates (RCC) and buffy coat were similar in both methods. However, the residual leukocytes and platelets in the RCC from the top‐and‐bottom system were significantly lower than in the conventional system, 140 ± 59 times 106 (mean ± SD) versus 762 ± 228 times 106, respectively (p<0.01). Both types of LPPC contained less than 107 leukocytes and could be stored for 7 days maintaining a pH above 6.5. We conclude that the top‐and‐bottom system enables automated and standardized preparation of RCC and plasma with a significantly better buffy‐coat removal than with manual processing.

Quality of Red Cell Concentrates in Relation to the Volume of the Buffy Coat Removed by Automated Processing in a Top and Bottom System

Abstract. The effect of automated removal of increasing volumes of buffy coat in a ‘top and bottom’ system on the composition of red cell concentrates (RCC) was investigated. The volume of the buffy coat was adjusted to group 1: 50 ml (n = 31), group 2: 70 ml (n = 31) and group 3:100 ml (n = 31), respectively. The numbers of platelets and leukocytes in the buffy coats were comparable between the groups, whereas the red cell volumes in the buffy coats showed a significant difference (17 ± 3.6 ml group 1, versus 22 ± 4.1 ml group 2 and 26 ± 3.8 ml group 3; p < 0.001). The volumes, hematocrits and cell counts of the RCC were not significantly different. The plasma volumes were inversely correlated with the volume of buffy coat removed, i.e. 268 ± 19 ml group 1, versus 257 ± 15 ml group 2 and 233 ± 20 ml group 3 (p<0.001). We conclude that in the ‘top and bottom’ system an increase of the volume of the buffy coat from 50 to 100 ml did not improve the quality of the RCC regarding contamination with leukocytes and platelets.

A new cellulose acetate filter to remove leukocytes from buffy-coat-poor red cell concentrates

Abstract. Transfusion of leukocyte‐free red cell concentrates (RCC) prevents or delays HLA immunization in multitransfused patients. We investigated a new cellulose acetate filter which was recently introduced to remove leukocytes from buffy‐coat‐poor RCC. It was found that the filtration time was only 10 min with buffy‐coat‐poor RCC in saline‐adenine‐glucose‐mannitol (SAG M; n=23), hematocrit being 62 ± 2% (SD). The red cell loss was 13.5 ± 2.6% and leukocyte removal was more than 99%. Routine filtration in SAG M (n=179) showed again that more than 99% of the leukocytes were removed from buffy‐coat‐poor RCC with an original leukocyte content of 804 ± (SD)458times 106. The red cell loss (12 ± 8.6%) was not diminished by increasing the amount of saline (0.9% NaCl) from 100 to 300 ml in an attempt to remove the retained red cells from the filter. We conclude that the new filter is reliable in rapidly removing more than 99% of the leukocytes from buffy‐coat‐poor RCC in SAG M solution.