How to Culture Plasmodium falciparum Field Isolates: Step-by-Step Protocol

From Patient to Petri Dish: Culturing Plasmodium falciparum from Field Isolates

In 1976, William Trager and James Jensen managed something malaria researchers had chased for decades. They kept Plasmodium falciparum alive outside the human body, growing continuously in nothing more exotic than a flask of red blood cells and culture medium. It sounds almost too simple. It wasn't.

Fifty years later, that same basic system, blood cells, medium, a warm incubator, still underpins how we study malaria in the lab. But there's a meaningful difference between growing a laboratory-adapted strain like 3D7 and coaxing a parasite fresh out of a patient's bloodstream into stable, continuous culture. Lab strains have already adapted to artificial conditions. Field isolates haven't. They arrive stressed, genetically diverse, and often reluctant to grow.

Here's what that process actually looks like, from the moment blood leaves a patient to the point where a flask of parasites is thriving on its own.

Why Field Isolates Are Different

A laboratory clone has been in culture for years, sometimes decades. It's been selected, whether researchers intended it or not, for the traits that make it easy to grow: tolerance of artificial media, resilience to handling, a forgiving growth rate.

A field isolate has none of that history. It comes straight from a patient, carrying whatever genetic background it evolved under in the wild, and it has to survive both the trauma of collection and the abrupt shift from a human host to a plastic flask. Some isolates adapt within days. Others take weeks. Some never do.

This is why timing matters so much at the start. Blood should be collected before antimalarial treatment begins, because drugs in the patient's system will still be acting on the parasites in vitro, and the sample needs to be processed and into culture as quickly as possible.

Collecting and Preparing the Sample

The process starts with a straightforward venous draw, typically a few milliliters into a tube containing an anticoagulant like ACD-A or CPD-A. From there, the sample is split.

One portion goes toward culture. The other, a smaller aliquot, is frozen immediately as a stabilate, essentially an insurance policy. If the culture fails to establish, or if something goes wrong downstream, the frozen stabilate preserves that original isolate for a second attempt later.

The culture-bound portion gets diluted substantially, roughly twenty-fold, in complete culture medium. Diluting this heavily isn't arbitrary. It brings the cell concentration down to something the culture system can actually support, and it happens as soon as possible after collection, because parasite viability starts declining the moment blood leaves the body.

Setting Up the Culture

Once diluted, the isolate goes into flasks under conditions that mimic, as closely as a plastic vessel can, the environment inside a human blood vessel: 37°C, a gas mixture low in oxygen and enriched in carbon dioxide, and a growth medium supplemented with serum-replacement additives like Albumax.

From here the culture is maintained by daily medium changes. Spent medium is aspirated gently, so as not to disturb the settled red cells, and replaced with fresh, pre-warmed medium. The flask is gassed briefly, sealed, and returned to the incubator. Do this every day, at roughly the same time if possible, and a viable isolate will slowly establish itself.

Parasitaemia, the proportion of red cells actually infected, is checked regularly by preparing a thin blood smear, staining it with Giemsa, and counting under oil immersion. When it climbs into the 6 to 8 percent range, the culture is diluted back down to around 1 percent with fresh uninfected red cells. Go much lower than that and, oddly, the culture can stall rather than thrive. There's a floor below which dilution does more harm than good.

Contamination is the other constant threat. A flask that's cloudy, foul-smelling, or simply behaving oddly should be discarded without hesitation. Aseptic technique, correctly, and every time, is the only thing standing between a healthy culture and a ruined one.

Getting the Stages to Line Up

Left alone, a culture will contain parasites at every stage of the 48-hour asexual cycle simultaneously, rings, trophozoites, and schizonts all mixed together. For many experiments that's a problem, because you need parasites at a known, uniform stage.

Sorbitol synchronization solves this by exploiting a quirk of parasite biology: infected red cells become more permeable as the parasite matures, so exposing a culture to a sorbitol solution selectively lyses the later-stage cells and leaves the young rings untouched. Run it once and you've narrowed the population. Run it again, roughly one cycle later, and the synchrony tightens further, since by then the survivors have all matured into rings together too.

Percoll gradient centrifugation takes a different approach, using density rather than membrane permeability to physically separate mature, hemozoin-laden schizonts from the younger stages, which is particularly useful when you need to enrich for schizonts specifically rather than simply narrow the age range.

Banking the Isolate

A field isolate that survives long enough to establish is worth preserving, both for future experiments and because recollecting from the same patient usually isn't possible.

Freezing follows a fairly gentle protocol built around glycerol as a cryoprotectant, added slowly and at room temperature to give it time to penetrate the cells before the drop to liquid nitrogen. Thawing reverses the process in careful stages, using a series of buffered solutions added dropwise to avoid osmotic shock, which would otherwise rupture the very cells you're trying to recover.

Done correctly, a stabilate frozen this way can sit in liquid nitrogen for years and still be revived into a thriving culture.

What Makes It Worth the Trouble

Culturing lab-adapted clones is, by comparison, almost easy. Culturing field isolates is slower, less predictable, and demands more attention at every step. But it's also the only way to study parasites as they actually exist in circulation, carrying whatever drug-resistance mutations, growth characteristics, or genetic diversity they picked up in the population they came from.

That's the whole point. A lab strain can tell you how P. falciparum behaves in principle. A field isolate tells you how it's behaving right now, in the patients walking into a clinic today.

Workflow Overview

Culturing Plasmodium falciparum field isolates can be divided into eight stages.

1. Prepare incomplete RPMI 1640.
2. Prepare complete RPMI 1640.
3. Prepare supporting reagents.
4. Process donor red blood cells.
5. Screen donor blood for contamination.
6. Establish parasite cultures.
7. Perform daily culture maintenance.
8. Monitor parasite growth and dilute cultures when required.

Each of these stages is described in detail below.

Before You Begin 

Successful parasite culture depends as much on preparation as it does on technique. Before beginning, ensure that all media have been prepared and filter sterilised, the incubator has reached 37°C, and all tissue culture flasks and complete RPMI have been pre-warmed. Maintaining sterile technique throughout the procedure is essential because bacterial or fungal contamination can rapidly destroy a parasite culture.

Field isolates are often less tolerant of environmental stress than long-established laboratory strains. Consequently, minimise the amount of time cultures spend outside the incubator and handle culture flasks gently to avoid disturbing the sedimented erythrocytes.

🧪 Adwoa Biotech Insight

One of the simplest ways to improve parasite recovery is to organise your workspace before removing cultures from the incubator. Arrange your pipettes, sterile tips, microscope slides and fresh medium in advance so that routine maintenance can be completed quickly while the parasites remain close to their optimal growth temperature.

Materials and Equipment 

Reagents 

• RPMI 1640 powder (Gibco)

• Albumax

• HEPES

• Dextrose

• Hypoxanthine

• Sodium bicarbonate

• Sodium hydroxide (NaOH)

• Hydrochloric acid (HCl)

• Distilled water

• Sodium chloride

• Sorbitol

• Glycerol

• Methanol

• Giemsa stain

• Giemsa buffer

• Immersion oil

Equipment 

• Class II Biological Safety Cabinet

• Magnetic stirrer

• pH meter

• Refrigerated centrifuge

• 0.2 μm filter unit

• 25 cm² tissue culture flasks

• 75 cm² tissue culture flasks

• Incubator maintained at 37°C

• Light microscope with a 100× oil immersion objective

• Micropipettes and sterile pipette tips

• Sterile Pasteur pipettes

• Glass microscope slides

Preparing Incomplete RPMI 1640

Incomplete RPMI 1640 forms the foundation of the parasite culture medium. It supplies amino acids, vitamins, inorganic salts and nutrients required for parasite growth while allowing additional supplements to be added during preparation of the complete culture medium. Because every culture established in the laboratory depends on this solution, careful preparation is essential.

Reagents 

Reagent	Quantity
RPMI 1640 powder (Gibco)	10.4 g (1 packet)
HEPES	7.15 g
Dextrose	2 g
Hypoxanthine (25 mg/mL in NaOH)	2 mL
Distilled water	To 1 litre

Procedure 

1. Add approximately 850 mL of distilled water to a large sterile conical flask.
2. Place the flask on a magnetic stirrer and slowly dissolve one packet (10.4 g) of RPMI 1640 powder.
3. Add 7.15 g HEPES and continue stirring until dissolved.
4. Add 2 g dextrose.
5. Add 2 mL hypoxanthine (25 mg/mL prepared in NaOH).
6. Continue stirring until the solution becomes completely clear.
7. Adjust the pH to 7.2 using sodium hydroxide or hydrochloric acid.
8. Top up the solution with distilled water to a final volume of 1 litre.
9. Filter sterilise through a 0.2 μm membrane filter inside a biological safety cabinet.
10. Store the medium at 4°C for up to two months.

🧪 Adwoa Biotech Insight

Each component of incomplete RPMI serves a specific purpose. HEPES helps maintain a stable pH while cultures are handled outside the incubator, dextrose provides an additional source of glucose for parasite metabolism, and hypoxanthine supplies purines that Plasmodium falciparum cannot synthesise itself. Preparing the medium accurately and maintaining the correct pH are essential for achieving consistent parasite growth.

Complete RPMI-1640 Preparation for Malaria Culture

A Step-by-Step Guide to Plasmodium Culture Media

If you’ve spent any time in a malaria lab, you know that Plasmodium falciparum is a picky eater. 

The goal here is to create a stable, nutrient-rich environment that mimics human physiological conditions while accounting for the common pitfalls of lab life; like the frustrating tendency of buffers to outgas or essential amino acids to degrade.

The media used (RPMI-1640) was developed at Roswell Park Memorial Institute in the 1960s. The "1640" refers to the formulation number assigned during its development, reflecting the extensive empirical (trial-and-error, experiment-based) optimisation that produced the medium.

RPMI-1640 has a fascinating history because it emerged during the period when mammalian cell culture was transitioning from empirical media recipes to more rationally designed formulations. For those like me, wondering what empirical means, it’s approaches that were based on observation, experimentation, and trial-and-error rather than a complete theoretical understanding. 

Origin of RPMI-1640

RPMI stands for: Roswell Park Memorial Institute

The medium was developed at the Roswell Park Comprehensive Cancer Center in Buffalo, New York.

The principal developers were:

• George E. Moore

• Robert E. Gerner

• Harold A. Franklin

during the 1960s.

The general components and amounts in 1L RPMI are: 

RPMI 1640: 10.44 g (The nutritional backbone, containing l-glutamine at a final conc. of 2mM). This is 5.22g if only making 500 mL

HEPES: 5.96 g (Your primary buffering agent). If making 500 mL, add 12.5 mL of 1M HEPES. If you buy the powdered RPMI, it likely comes with this already.

NaHCO3​: 58 mL of 3.6%  (final is 2g/L ; For pH stability and gas exchange). Sodium bicarbonate is notorious for outgassing (releasing CO2​), which can cause your pH to drift upward over time. To combat this, we add it just before use.

Hypoxanthine: 50 mg (200uM, Essential for parasite purine salvage)

Gentamicin: 20 ug/mL stock (Your antibiotic shield). For 1L you can add 20 mg of gentamicin

Ultrapure/sterile H2​O: 960 mL

1M NaOH: For dissolving the hypoxanthine

Concentrated HCL and NaOH: For final pH adjustment

Related Content

Ready to put this media to use? Our step-by-step Plasmodium falciparum culture protocol shows exactly how Complete RPMI-1640 fits into culture initiation and maintenance.

References:

1.     Lopez-Perez, M., & Seidu, Z. (2022). Establishing and Maintaining In Vitro Cultures of Asexual Blood Stages of Plasmodium falciparum. Methods in Molecular Biology. 

2.     Maier, A. G., & Rug, M. (2013). In vitro culturing Plasmodium falciparum erythrocytic stages. Methods in Molecular Biology. 

3.     Trager, W., & Jensen, J. B. (1976). Human malaria parasites in continuous culture. Science, 193(4254), 673–675. 

4.     World Health Organization. (2023). World Malaria Report 2023. Geneva: WHO.

How to Isolate Red Blood Cells for Plasmodium falciparum Culture: Protocol, History, and What Your Tube Is Actually Doing

 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. 

Fig. 1a: Peripheral blood smear prepared from a sample stored for approximately three weeks. Numerous red blood cells display evenly spaced spiky membrane projections characteristic of a phenomenon known as crenation (echinocyte formation). Normal RBCs appear as biconcave discs (like a doughnut without a hole) with smooth edges. The center looks slightly lighter.

Fig. 1b: Peripheral blood smear showing normal RBCs. Note the biconcave discs with smooth edges and a slightly light center.

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 (Fig. 2). 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. 

Fig. 2: A 2.5% Giemsa stained thin blood smear showing ring stage parasites.

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

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, research groups commonly use O+ and A+ erythrocytes due to their high global availability (approximately 75%); however, the parasite is capable of proliferating in any ABO blood type.

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 (Fig.1). 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.   

Fig. 3: A 2.5% Giemsa stained thin blood smear showing Trophozoites inside RBCs.

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.   

Does Male/Female Donor Matter? Addressing a Persistent Lab Myth

There are also myths that persist in lab culture. 

One of the more common ones is that blood from female donors is somehow less suitable for parasite culture. 

There is no strong evidence to support this. 

When differences are observed, they are almost always explained by cell quality, handling, or donor-specific properties, not male or female donor. 

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). We are especially grateful to our dedicated O+ blood donor (Derrick Sackitey), whose consistent and reliable donations have made our P. falciparum 3D7 cultures possible.

References

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