Skip to Content


January-April 2004
Fresh Frozen Plasma Is Not a Cellular Blood Component: Transfusion Practice in Bone Marrow and Peripheral Blood Stem Cell Transplant Recipients

By Aida Narvios, Geovanna Figueredo, Jeffrey J. Tarrand, Yang O. Huh, and Benjamin Lichtiger

Copyright 1993-2010 The University of Texas MD Anderson Cancer Center, Houston, Texas. All rights reserved.

Cellular blood components (red blood cells and platelets) are naturally contaminated with white blood cells (WBCs), which may induce a series of reactions, such as alloimmunization, graft-versus-host disease (GVHD), and transmission of infectious diseases, including cytomegalovirus (CMV), human T-lymphotropic virus 1, and Epstein-Barr virus (1). Fresh frozen plasma (FFP) and cryoprecipitates are usually considered to be acellular blood components or slightly contaminated with cellular elements too low in number to cause adverse events in which WBCs could be implicated (2). Currently, there is evidence in the literature suggesting that there are enough viable WBCs in FFP to induce GVHD in immunocompromised patients (2-4). One report even suggests that the presence of WBCs in FFP is a cause of posttransfusion GVHD (5).

Transmission of CMV via transfusion of red blood cells and platelets has been well documented (6-8). These blood components may play a significant role in the transmission of CMV via transfusion, especially in immunocompromised patients. Therefore, it has been suggested that leukoreduction of such blood components can prevent CMV infection (9,10). However, reports in the medical literature have provided clinical evidence that transfusion of FFP may not be an important factor in the transmission of CMV (11- 17). In light of these findings, the purpose of our study was to determine whether the number of WBCs present in units of FFP warrants leukoreduction of FFP prior to transfusion to prevent CMV infection.

Materials and Methods

Production of FFP

Fifty units of FFP were produced in routine fashion at The University of Texas MD Anderson Cancer Center Blood Bank in accordance with regulatory standards and in compliance with the standard operating procedures of the blood bank. Briefly, FFP was prepared from units of whole blood and frozen at -18°C within the required time frame for the anticoagulant or collection process (18,19).

Of the 50 units of FFP produced for this study, 20 were used for enumeration of leukocytes before and after filtration via a manual method with a Nageotte chamber. Ten units were used for determination of the WBC populations present after filtration via flow cytometry. Samples of the remaining 20 units were analyzed via cell culture to determine whether CMV was present.

Leukoreduction of FFP

Twenty units of FFP were filtered through standard leukoreduction red blood cell filters (Sepacell 500; Baxter, Deerfield, IL). There were no FFP-specific leukoreduction filters available in our blood bank. Leukoreduction was performed at room temperature without any priming of the filter. The filtration period for each unit of FFP with an average volume of 220 ml was 5-10 minutes. Each filter was used for filtration of four units of FFP.

Counting of WBCs

WBCs from the 20 units of FFP samples were counted before and after filtration. This was done manually with the aid of a Nageotte chamber. Turk's solution was added to the samples for visualization of the nuclei of the WBCs with a standard microscope with a 10x and 40x ocular as described previously (20).

Flow cytometry

Ten units of unfiltered FFP were analyzed via flow cytometry after being counted with a Z1 Series Coulter Counter (Beckman Coulter, Miami, FL). Immunophenotypic analysis was conducted with the use of a FACScan (BD Biosciences, San Jose, CA) with different antibodies, such as Leu4 (CD3), Leu3 (CD4), Leu2 (CD8), Leu12 (CD19), Leu1 (CD5), LeuM1 (CD15), CD41, CD13, CD33, CD45, and LeuM3 (CD14).

The samples were incubated with 10 microliters of each antibody at 20-80°C for 15 minutes in the dark and washed twice with a solution of buffered phosphate. The samples were then resuspended, fixed in a 1% solution of paraformaldehyde, and analyzed with the CellQuest software program (BD Biosciences). Lymphocytes, monocytes, and granulocytes were discriminated based on the density of CD45 and characteristic pattern of scattering on the cell surface.

Culture and antigen detection of CMV in FFP

One sample from each of the 20 units of unfiltered FFP was analyzed to determine the presence of CMV. This was performed via direct fluorescent antigen detection with the use of a monoclonal antibody directed against the CMVpp65 structural protein and culture-based methods. Briefly, FFP was thawed rapidly in a 35°C water bath, and the cellular component was recovered via centrifugation at 200g for 10 minutes at 40°C. Antigen detection was performed with a CMVpp65 antigenemia assay kit (Chemicon International, Temecula, CA). Appropriate positive and negative controls were used. The cells were washed in phosphate-buffered saline, and erythrocytes were lysed with cold ammonium chloride. The typical yield of 450 ml of FFP was 1 x 109 cells. Slides were stained and interpreted as per antigen detection of CMV. For the culture-based methods, 2 ml of residual supernatant was used to inoculate two MRC5 shell vials (0.5 ml of inoculum each) and two diploid human foreskin fibroblast cells lines (SF cells inoculated with 0.5 ml each). The shell vials were incubated for 18-24 hours and read with the use of the Bartels CMV stain (Chemicon International, Temecula, CA), and the SF cells were interpreted for cytopathic effect for 21 days (21).

Table 1. Enumeration of WBC Counts with the Nageotte Chamber*

Sample no.Unit weight (g)Prefiltration WBCs (x 106)Residual WBCs (x 106)Leukoreduction (%)

*SD, standard deviation.


Table 2. Results of Flow Cytometry Analysis After Filtration* (PDF)

*No., sample number; Vol., volume; WBC, white blood cell count; Plt., platelet count; Lym., lymphocytes; Mon., monocytes; Gra., granulocytes; Conc., concentration.


Table 1 shows the mean WBC counts before and after filtration of FFP as determined manually with the Nageotte chamber. The mean number of WBCs in the prefiltration samples was 1.57 x 106 (range, 0.11-3.25 x 106). This was below the number required by regulatory standards for leukoreduced products but was clinically significant enough for our bone marrow transplant recipients receiving CMV- unscreened blood products. The mean number of residual WBCs after filtration was 0.00246 x 106 (range, 0.001-0.010 x 106). In addition, the mean percentage removal of WBCs through leukoreduction was 98.52% (range, 96.75%-99.99%). This showed highly efficient removal of WBCs.

Table 2 shows the results of our postfiltration sample analysis via flow cytometry. The mean number of residual WBCs after leukoreduction was 0.19 x 106 (range, 0.03-0.29 x 106 ). In the 10 leukoreduced FFP units, we found that the residual WBCs were composed of a mean of 10.9% lymphocytes (range, 2%-25%), 3.8% granulocytes (range, 1%-10%), and 5% monocytes (range, 1%-10%). Flow cytometric analysis showed that the mean concentration of CD4, CD8, and CD19 was 4.2%, 2.2%, and 2.7%, respectively. Dead cells were also seen, which could have accounted for the remaining cells. Furthermore, the mean number of platelets was 16.57 x 106 (range, 14.0-21.7 x 106). There was no evidence of CMV in any of the samples.


Several authors have reported finding large numbers of WBCs in units of FFP. For example, Willis et al. (4) reported a contamination rate of more than 1 x 106 WBCs per unit of FFP. Similarly, Bernvil et al. (2) reported a mean of 2 x 106 WBCs per unit of FFP, and Wieding et al. (3) found a mean of 3.2 x 106 WBCs per unit.

The findings of the present study are quite similar to those reported by the above-mentioned authors. Furthermore, we were able to quantitate the platelets in the FFP units. Although the number of platelets in FFP appears to be low, their presence may play an important role in the development of platelet refractoriness. Lymphocytes present in FFP may also play a role in the development of posttransfusion GVHD. Another important finding in our study was that the presence of CMV could not be ascertained despite the use of various methods of detecting it.


The results of this study demonstrate that FFP is not acellular but is contaminated with a substantial number of WBCs and platelets. The presence of these cells may lead to serious adverse effects in susceptible immunocompromised hosts who receive a transfusion of FFP. In light of these findings, bone marrow and peripheral blood stem cell transplant recipients who are supposed to receive leukoreduced cellular blood components could benefit from transfusion of FFP that has been subjected to the manipulation described herein. At MD Anderson Cancer Center, units of FFP that are destined for transfusion in patients who are on a leukoreduction protocol, including bone marrow and peripheral blood stem cell transplant recipients, are administered after filtration with a red blood cell leukoreduction filter (four units of FFP per filter). This is performed at the patient's bedside. Although we are aware that plasma filters are commercially available, we found the red blood cell leukoreduction filter to be economical and practical for this purpose.


We thank Dr. Elpidio Pena for assisting us in this study.


  1. Mintz PD. Quality assessment and improvement of transfusion practices. Hematol Oncol Clin North Am 9:219-232, 1995.
  2. Bernvil S, Abdulatiff M, Al-Sedairy S, Sasich F, Sheth K. Fresh frozen plasma contains viable progenitor cells--should we irradiate? Vox Sang 67:405, 1994.
  3. Wieding JU, Vehmeyer K, Dittman J, et al. Contamination of fresh frozen plasma with viable white cells and proliferative stem cells (letter). Transfusion 34:185-186, 1994.
  4. Willis J, Lown J, Simpson M, Erber W. White cells in fresh frozen plasma: Evaluation of a new white cell-reduction filter. Transfusion 38:645-649, 1998.
  5. Rubinstein A, Rad J, Cottier H, et al. Unusual combined immunodeficiency syndrome exhibiting kappa-IgD paraproteinemia, residual gut-immunity and graft vs host reaction after plasma infusion. Acta Paediatr Scand 62:365-372, 1973.
  6. Adler SP. Transfusion-associated CMV infection. Rev Infect Dis 5:977-993, 1983.
  7. Schrier RD, Nelson JA, Oldstone MBA. Detection of human CMV in peripheral blood lymphocytes in a natural infection. Science 230:1048- 1051, 1985.
  8. Stanier P, Taylor DL, Kitchen AD, et al. Persistence of CMV in mononuclear cells in peripheral blood from blood donors. Br Med J 299:897-898, 1989.
  9. Andreu G. Role of leukocyte depletion in the prevention of transfusion-induced CMV infection. Semin Hematol 28:26-31, 1991.
  10. Bowden RA, Slichter SJ, Sayers MH, et al. Use of leukocyte-depleted platelets and CMV infection after marrow transplant. Blood 78:246-250, 1991.
  11. Yeager AS, Grumet FC, Hafleigh EB et al. Prevention of transfusion- acquired CMV infections in newborn infants. J Pediatr 98:281-287, 1981.
  12. Demmler GJ, Brady MT, Bijou H, et al. Posttransfusion cytomegalovirus infection in neonates: Role of saline-washed red blood cells. J Pediatr 108:762-765, 1986.
  13. Brady MT, Milam JD, Anderson DC. Use of deglycerolized red blood cells to prevent posttransfusion infection with CMV in neonates. J Infect Dis 150:334-339, 1984.
  14. Components from Whole Blood Donations. AABB Technical Manual. 14th Edition. American Association of Blood Banks. Bethesda, MD, 2002, pp. 165-166.
  15. Tegtmeier GE. Cytomegalovirus and blood transfusion. In: Dod RY, Barker LF (eds), Infection, Immunity and Blood Transfusions. Proceedings of the 16th Annual Scientific Symposium of the American Red Cross. Washington, DC, 1984, volume 182, pp. 175-199.
  16. Cheeseman SH, Sullivan JL, Brettler DB, Levine PH. Analysis of cytomegalovirus and Epstein-Barr virus antibody response in treated hemophiliacs: implications for the study of acquired immune deficiency syndrome. JAMA 253:83-85, 1984.
  17. Landay A, Poon MC, Abo T, et al. Immunologic studies in asymptomatic hemophilia patients: Relationship to acquired immune deficiency syndrome (AIDS). Clin Invest 71:1500-1504, 1983.
  18. Gorlin JB (ed), Standards for Blood Banl and Transfusion Services. 21st ed. American Association of Blood Banks. Bethesda, MD, 2002.
  19. American Association of Blood Banks, American National Red Cross, and America's Blood Centers. Circular of Information for the Use of Human Blood and Blood Components. American Association of Blood Banks. Bethesda, MD, 2002.
  20. Enck RE, Betts RF, Brown MR, Miller G. Viral serology (hepatitis B virus, cytomegalovirus, Epstein-Barr virus) and abnormal liver function tests in transfused patients with hereditary hemorrhagic diseases. Transfusion 19:32-38, 1979.
  21. Bowden R, Sayers M. The risk of transmitting CMV infection by fresh frozen plasma. Transfusion 30:762-763, 1990.

© 2014 The University of Texas MD Anderson Cancer Center