RBC Isolation for Plasmodium falciparum Culture

Isolating red blood cells from whole blood sounds like a simple procedural step. Spin it down, remove the plasma, wash a few times, done. But every step of this process has a reason rooted in biology, chemistry, and in some cases, history that stretches back over a century.

Today, we are going to walk through the full RBC isolation protocol for P. falciparum culture, explain the biology and chemistry behind every step, and take a detour into the remarkable history of the people and discoveries that gave us the tools we use every day.

Why Do We Isolate RBCs in the First Place?

Whole blood is not just red blood cells. It is a mixture of plasma, white blood cells, platelets, and those red blood cells all suspended together. For P. falciparum culture, most of what is in whole blood is either useless or actively harmful.

Plasma carries antibodies and complement proteins that can kill parasitised cells. White blood cells recognise and destroy infected RBCs as part of normal immune function. Platelets aggregate and introduce background interference in assays. None of these belong in your culture flask.

By isolating and washing the RBCs, you strip all of that away and give the parasite exactly what it needs: clean, metabolically competent red blood cells in a defined, controlled environment where invasion, replication, and stage progression can happen reliably and reproducibly.

Step One: Choosing the Right Anticoagulant and the History Behind it

Before you collect a single drop of blood, you need to make a decision that will shape everything that follows: which anticoagulant tube do you use?

For P. falciparum culture, the answer is ACD : acid, citrate dextrose. But to understand why ACD is the right choice and why the alternatives fall short, it helps to understand what is actually inside that yellow-capped tube. And to understand that, we need to go back over 100 years.

1914 to 1918: The Problem of Clotting Blood

In the early 20th century, blood transfusion was possible in theory but chaotic in practice. Blood clotted within minutes of leaving the body, which meant donor and recipient had to be physically connected during a transfusion - hardly practical on a battlefield.

In 1914, researchers in Belgium, Argentina, and the United States independently discovered that sodium citrate could prevent clotting by removing free calcium ions from the blood. Calcium is an essential cofactor for the coagulation cascade, so binding it halts clotting entirely. By 1915, Richard Lewisohn at Mount Sinai Hospital had refined this into a practical anticoagulant method, and Richard Weil had shown that citrated blood could be refrigerated for short periods. The basic calcium chelation principle sitting inside an ACD tube today was first worked out more than 110 years ago.

Oswald Hope Robertson took this further during World War I, adding dextrose to the citrate solution to feed the RBCs during storage and extending their usable life.

1943: Loutit, Mollison, and the ACD Formulation

In 1943, British researchers J.F. Loutit and Patrick Mollison solved a persistent problem with earlier citrate-glucose solutions: the glucose would caramelise during heat sterilisation of the tubes, making the solution unusable. Their breakthrough was to add citric acid to lower the pH, which prevented caramelisation and also impaired residual thrombin activation, adding a secondary anticoagulant effect. This three-component formulation - citric acid, sodium citrate, and dextrose, became ACD solution.

Each component has a distinct job. Sodium citrate chelates free calcium ions to prevent clotting. Dextrose serves as an energy source for RBCs during storage, maintaining their metabolic function and extending viability. Citric acid lowers the pH to prevent the dextrose from caramelising during sterilisation and provides a secondary anticoagulant effect by inhibiting residual thrombin.

ACD enabled refrigerated storage of whole blood and was formally adopted by blood banks at the end of World War II. It has remained the preferred anticoagulant for research blood applications, including P. falciparum culture, ever since.

One limitation of ACD is worth knowing: the acidic pH does not maintain 2,3-bisphosphoglycerate (2,3-BPG) levels well during storage. 2,3-BPG is a molecule that helps haemoglobin release oxygen to tissues. This drove the development of CPD (citrate-phosphate-dextrose) in 1957, with phosphate added to buffer the pH, and then CPDA-1, which extended RBC shelf life to 35 days. For P. falciparum culture, where oxygen delivery to tissues is irrelevant, ACD remains the standard.

Why Not Heparin? Why Not EDTA?

Heparin is common in clinical settings and might seem like a perfectly reasonable substitute. It is not; at least not for P. falciparum culture.

Heparin inhibits merozoite invasion of RBCs. That is the very process you are trying to study. Some labs deliberately use this property to synchronise cultures or study the invasion mechanism in isolation, but for routine culture it is actively counterproductive. Using heparin-collected blood is essentially adding a parasite invasion blocker to your experiment without meaning to.

EDTA is widely used in haematology and molecular work but leads to faster RBC degradation compared to ACD and is not suitable for routine culture applications. Stick with ACD.

Step Two: What to Do the Moment That Tube Is Full

The moment blood is drawn into your ACD tube, the clock starts. The manufacturer's instructions are clear: invert the tube 8 to 10 times immediately. Gently. No shaking. Shaking causes foaming and haemolysis, which is the last thing you want before you have even started processing.

The reason for the inversion is straightforward: you need the ACD anticoagulant in contact with all of the blood before clotting begins. Inadequate mixing means you risk microclots forming in the sample, which will compromise your yield and your cell quality.

After mixing, get the tube to the centrifuge as soon as practically possible and always within four hours of collection. At room temperature, ongoing metabolic activity in the whole blood - from both RBCs and white blood cells - begins degrading sample quality beyond this window. The ACD preserves the cells during storage, but it does not stop metabolic processes entirely. The four-hour window is not arbitrary; it is the point at which that degradation starts becoming significant.

Step Three: Centrifugation and the Three Layers

Centrifuge whole blood at 400 to 500 x g (gravity) for 5 to 10 minutes. This separates the blood into three distinct layers that you need to be able to identify clearly before you proceed.

At the top, you will see the plasma layer : pale yellow, sometimes slightly cloudy. Beneath that is the buffy coat, a thin whitish layer containing the white blood cells and platelets. At the bottom is the packed red blood cell pellet: dark red and compact.

All three layers are easy to distinguish once you have seen them a few times, but the buffy coat is the one that demands your full attention.

Step Four: Remove the Plasma

Carefully aspirate and discard the plasma layer. Plasma contains antibodies, complement proteins, and a range of undefined factors that interfere with culture reproducibility.

Step Five: The Buffy Coat

This is a critical manual step in the protocol, and it is the one most likely to cause problems if you are not deliberate about it.

The buffy coat sits directly on top of the RBC pellet. It must be removed completely. Even a small number of residual leukocytes will recognise and destroy infected RBCs, interfering with your parasite growth in ways that can look like a culture problem when it is actually a preparation problem.

Rotate the tube slowly as you aspirate, working your way around the full circumference of the interface. Take your time. It is better to sacrifice a small number of RBCs at the top of the pellet than to leave leukocytes behind.

Step Six: Three Washes in Incomplete RPMI

Add incomplete RPMI 1640 in a volume three to five times that of the RBC pellet, resuspend gently, centrifuge at 400 to 500 x g for 5 minutes, and discard the supernatant. Repeat three times in total.

Most protocol guides describe this as removing residual plasma proteins and any remaining leukocytes. That is true, but it undersells what the washes are actually doing. To understand the full picture, you need to go back to the ACD chemistry we discussed earlier.

Remember that ACD works by chelating ionised calcium. The citrate molecules bind calcium so effectively that they halt the entire coagulation cascade. That is exactly what you want during collection and storage. But calcium in your culture media is not a problem. In fact, calcium is a requirement. Calcium-dependent processes are involved in the RBC membrane dynamics that P. falciparum merozoites exploit during invasion. Residual citrate carried over from the ACD solution into your culture flask will continue chelating free calcium in the RPMI, potentially interfering with the very invasion process your experiment depends on.

The three washes are what allow you to have both. We use ACD for its protective properties during collection and storage, and you wash it away before the cells enter our culture environment. Each wash dilutes and removes more residual citrate alongside the plasma proteins and leukocyte debris.

The Hidden Variable: RPMI Temperature During Washing

Most standard blood processing protocols call for washing RBCs in phosphate-buffered saline (PBS), typically at room temperature. P. falciparum culture protocols typically use incomplete RPMI 1640 instead of PBS for washing, but what is the best temperature for the washing? Should that RPMI be cold, room temperature, or pre-warmed?

Our lab uses cold incomplete RPMI 1640 straight from the refrigerator for washing steps. This is practical, efficient, and has worked reliably for our cultures. But the literature suggests we might be leaving performance on the table.

The WWARN (WorldWide Antimalarial Resistance Network) protocol specifically recommends warming RPMI medium to 37°C in water-bath or heater block (19).

The biological rationale centers on membrane stability during washing. Human RBCs show no hemolysis at temperatures at or below 37°C during extended incubation, but cold processing temperatures can increase hemolysis, particularly with repeated washing steps (7).

Step Seven: Resuspend at 50% Haematocrit and How Long Can You Keep Washed RBCs?

Add complete RPMI 1640 supplemented with Albumax or human serum, in a 1:1 ratio with the packed RBC pellet. This creates a 50% haematocrit working stock that can be stored at 4 degrees C.

Here is something worth knowing about storage duration: washed RBCs stored at 50% haematocrit in complete RPMI can technically support P. falciparum culture for up to four weeks. But the critical word there is technically. Published data show that growth rates decline progressively with storage length, falling by approximately 62% after four weeks compared to freshly washed cells (4). The decline becomes significant at the two-week mark. In practice, we aim to use washed RBCs within two weeks for reliable, reproducible results. Between two and four weeks they will still work, but you should expect reduced growth rates and account for this in your experimental interpretation.

For gametocyte production specifically, at least one published protocol recommends using RBCs less than one week old, stored in complete medium with human serum. Gametocytogenesis is sensitive to RBC quality in ways that asexual blood stage culture is not.

One rule that is absolute and non-negotiable: never freeze RBCs. Ice crystal formation during freezing destroys the cell membrane irreversibly. There is no recovering from it.

Step Eight: Dilute to 5% Haematocrit for Culture

P. falciparum is typically cultured at no more than 5% haematocrit. Dilute your 50% working stock 1:10 in complete RPMI to achieve this.

A Critical Note on Donor Compatibility: O+ Is Necessary, Not Sufficient

In 1912, Roger Lee and Paul Dudley White coined the term universal donor to describe O blood type, demonstrating that O blood, which lacks the A and B antigens that trigger immune rejection, could be given to patients of any blood group. For emergency transfusion, O negative is the true universal donor because it also lacks the Rh antigen. For P. falciparum culture, O positive is the convention: the Rh factor is irrelevant to parasite invasion, and O positive is far more common, making it far more practical for research blood banking.

We rely on a single validated O+ donor for our routine P. falciparum 3D7 cultures. When we tested a second O+ donor ; same blood type, same protocol, same everything, the parasites failed to invade entirely. The cultures did not grow at all.

This is not an isolated finding. Published research confirms that parasite invasion and replication vary substantially between donors even under otherwise identical conditions, and that these differences reflect stable, intrinsic properties of the RBC. Invasion efficiency does not correlate reliably with blood group or surface receptor levels. The explanation lies in RBC membrane biomechanics: membrane tension, deformability, and cytoskeletal protein composition all vary between individuals, and there is a membrane tension threshold above which merozoite invasion simply cannot proceed. These properties are invisible to standard ABO and Rh blood typing.

What this means practically is straightforward. When you find a donor whose blood reliably supports your culture, validate them thoroughly, document them, and hold onto them. If your donor becomes unavailable and you need to switch, do not assume any O+ blood will work. Test the new donor in parallel with your existing cultures before making a full switch.

A Practical Note on Delayed Processing:

Our standard practice is to wash ACD-stored whole blood within 24 to 48 hours of collection. In a perfect world, that is always possible. In a real laboratory, sometimes it is not.

On one occasion, we could not process the blood within our usual window. The tubes sat unprocessed in ACD at 4 degrees C for 96 hours before we were able to centrifuge and wash them. We were not sure what we would get.

The cultures grew. The parasites invaded. We were reassured. But we also observed that invasion rates appeared lower than we would typically expect.

What we can say with confidence is this: 96-hour cold-stored blood in ACD is not a dead end. Our observation suggests that 96-hour cold-stored ACD blood can serve as a workable contingency when normal processing is not possible. But it is a single observation with a confounding variable. It is not a formal validation, and we would not recommend routine delayed processing on the basis of it alone.

Beyond Malaria

RBC isolation is a foundational technique that extends well beyond P. falciparum work. It underpins transfusion medicine research on blood group antigens, haemoglobinopathy studies in sickle cell disease and thalassaemia, culture of other blood parasites including Babesia, flow cytometry controls in immunology, and membrane protein studies in cell biology.

Acknowledgements

This work was shaped by everyone who contributed to it - from the people who collected the blood (Prince Horlutu and Edward Dumashie), to those who processed it (Yakubu Osmanu and Eyram Adoboe), to those who support the work in the lab (Rita Afriyie and The Anita Ghansah laboratory).

References

BenchChem. (n.d.). In vitro culture and amodiaquine susceptibility testing of Plasmodium falciparum [Application note]. https://pdf.benchchem.com/18/Application_Notes_and_Protocols_In_Vitro_Culture_and_Amodiaquine_Susceptibility_Testing_of_Plasmodium_falciparum.pdf
Bennett-Guerrero, E., Veldman, T. H., Doctor, A., Telen, M. J., Ortel, T. L., Reid, T. S., Mulherin, M. A., Zhu, H., Buck, R. D., Califf, R. M., & McMahon, T. J. (2007). Evolution of adverse changes in stored RBCs. Proceedings of the National Academy of Sciences, 104(43), 17063–17068. https://doi.org/10.1073/pnas.0708160104
Clark, M. A., Goel, V. K., Cabrera, A., Clawson, M. L., Hellerstein, M. K., Mohandas, N., & Llinás, M. (2009). Host cell selection by Plasmodium falciparum is a determinant of gene expression. Nature, 458(7239), 703–706. https://doi.org/10.1038/nature07739
Ding, X. C., Ubben, D., & Kremsner, P. G. (2009). Plasmodium falciparum cultivation using the Petri dish: Revisiting the effect of the age of erythrocytes and the interval of medium change. Korean Journal of Parasitology, 47(2), 135–141. https://doi.org/10.3347/kjp.2009.47.2.135
Fischbach, A., Goetz, A., Karma, S., Bousema, T., & Sutherland, C. J. (2022). Homeostasis of extracellular ATP in uninfected RBCs from a Plasmodium falciparum culture and derived microparticles. Biochimica et Biophysica Acta (BBA) - Biomembranes, 1864(8), 183955. https://doi.org/10.1016/j.bbamem.2022.183955
Hess, J. R. (2010). Red cell storage. Journal of Proteomics, 73(3), 368–373. https://doi.org/10.1016/j.jprot.2009.10.006
Lippi, G., Blanckaert, N., Bonini, P., Green, S., Kitchen, S., Palicka, V., Vassault, A., & Plebani, M. (2008). Haemolysis: An overview of the leading cause of unsuitable specimens in clinical laboratories. Clinical Chemistry and Laboratory Medicine, 46(6), 764–772. https://doi.org/10.1515/CCLM.2008.170
Mata-Cantero, L., Lafuente, M. J., Sanz, L., & Rodríguez, M. S. (2014). Magnetic isolation of Plasmodium falciparum schizonts iRBCs to generate a high parasitaemia and synchronized in vitro culture. Malaria Journal, 13, Article 112. https://doi.org/10.1186/1475-2875-13-112
Mohandas, N., & Chasis, J. A. (1993). Red blood cell deformability, membrane material properties and shape: Regulation by transmembrane, skeletal and cytosolic proteins and lipids. Seminars in Hematology, 30(3), 171–192.
Nakao, M., Nakao, T., & Yamazoe, S. (1960). Adenosine triphosphate and maintenance of shape of the human red cells. Nature, 187, 945–946. https://doi.org/10.1038/187945a0
Pathak, A. K., Shiau, J. C., Thomas, M. B., & Murdock, C. C. (2018). Cryogenically preserved RBCs support gametocytogenesis of Plasmodium falciparum in vitro and gametogenesis in mosquitoes. Malaria Journal, 17, Article 457. https://doi.org/10.1186/s12936-018-2612-y
Radfar, A., Méndez, D., Moneriz, C., Linares, M., Marín-García, P., Puyet, A., Diez, A., & Bautista, J. M. (2009). Synchronous culture of Plasmodium falciparum at high parasitemia levels. Nature Protocols, 4(12), 1899–1915. https://doi.org/10.1038/nprot.2009.198
Schuster, F. L. (2002). Cultivation of Plasmodium spp. Clinical Microbiology Reviews, 15(3), 355–364. https://doi.org/10.1128/CMR.15.3.355-364.2002
Silvestrini, F., Alano, P., & Williams, J. L. (2000). Commitment to the production of male and female gametocytes in the human malaria parasite Plasmodium falciparum. Parasitology, 121(5), 465–471. https://doi.org/10.1017/s0031182000006760
Sutera, S. P., & Mehrjardi, M. H. (1975). Deformation and fragmentation of human red blood cells in turbulent shear flow. Biophysical Journal, 15(1), 1–10. https://doi.org/10.1016/S0006-3495(75)85787-0
Suzuki, Y., Tateishi, N., Soutani, M., & Maeda, N. (1996). Deformation of erythrocytes in microvessels and glass capillaries: Effects of erythrocyte deformability. Microcirculation, 3(1), 49–57. https://doi.org/10.3109/10739689609146782
Trager, W., & Jensen, J. B. (1976). Human malaria parasites in continuous culture. Science, 193(4254), 673–675. https://doi.org/10.1126/science.781840
Weed, R. I., LaCelle, P. L., & Merrill, E. W. (1969). Metabolic dependence of red cell deformability. The Journal of Clinical Investigation, 48(5), 795–809. https://doi.org/10.1172/JCI106038
WorldWide Antimalarial Resistance Network (WWARN). (2013). Culture of Plasmodium falciparum blood stages (Version 1.0). https://www.iddo.org/sites/default/files/publication/2023-09/inv01-culture-of-plasmodium-falciparum-blood-stages.pdf
Zimrin, A. B., & Hess, J. R. (2009). Current issues relating to the transfusion of stored red blood cells. Vox Sanguinis, 96(2), 93–103. https://doi.org/10.1111/j.1423-0410.2008.01117.x

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