Transporting Malaria Blood Samples: Temperature Matters

 

I still remember the first time a batch of field-collected blood arrived at the lab looking perfectly fine, and turned out to be completely useless. Nobody had told the courier that malaria parasites care, quite a lot, about temperature. So let's fix that gap. If you're planning to collect P. falciparum infected blood in the field, the very first decision you need to make isn't about equipment or anticoagulant. It's about what you're trying to preserve, because asexual parasites and gametocytes want opposite conditions.

If you want asexual ring stages: keep it cold

Collect into ACD or CPD vacutainers rather than heparin, which can interfere with downstream processing and invasion assays (Thiam et al., 2022). Then refrigerate the tubes at 4°C, or pack them in a cooler with cold packs, making sure the tubes never touch the ice directly. Direct contact freezes the red cells locally and kills the parasites instantly. You have a window here, but it's not a generous one. Aim to process within 2 to 6 hours, since viability drops off once whole blood sits at 4°C for more than a day or two (Okitwi et al., 2022).

If you want gametocytes: never refrigerate

This one trips people up because it goes against instinct. Cold blood feels like careful, responsible blood. But cooling activates gametocytes prematurely and wrecks their infectivity before they ever reach a mosquito. Instead, keep the sample near body temperature, somewhere around 35.5°C, using something as simple as a thermos flask filled with warm water. Soumare and colleagues (2021) tested exactly this approach across sites in Burkina Faso and The Gambia and found that gametocyte infectivity held up for up to four hours stored this way. Four hours is your practical ceiling, so plan your transport route with that in mind.

If the lab is more than a day away: cryopreserve on site

Sometimes neither option above is realistic. If you're looking at more than 24 hours before the sample reaches a facility, don't transport it raw at all. Centrifuge in the field to remove the plasma, add a glycerol-based cryoprotectant like Glycerolyte 57 dropwise to the packed red cell pellet, then move the aliquoted vials into a portable liquid nitrogen dry shipper or dry ice. Stanisic et al. (2015) used cryopreservation to build clinical-grade P. falciparum cell banks this way, and the same principle scales down to a single field sample.

The thread running through all three options is the same: decide what you need before you collect, not after. A tube of blood that's been babied at exactly the wrong temperature is no better than one left in the sun.

Related Content

ACD-A Blood Collection Tubes for P. falciparum Culture: https://adwoabiotech.blogspot.com/2026/07/acd-blood-collection-tubes-for-p.html

RBC Isolation for Plasmodium falciparum culture: https://adwoabiotech.blogspot.com/2026/04/rbc-isolation-plasmodium-falciparum-culture%20.html

Calculating Haematocrit when Passaging Plasmodium falciparum Cultures: https://adwoabiotech.blogspot.com/2025/06/how-to-calculate-hematocrit-for-malaria.html

References

Okitwi, M., Orena, S., Katairo, T., Tumwebaze, P. K., Byaruhanga, O., Nsobya, S. L., Conrad, M. D., Bayles, B. R., Rosenthal, P. J., & Cooper, R. A. (2022). Impact of short-term storage on ex vivo antimalarial susceptibilities of fresh Ugandan Plasmodium falciparum isolates. Antimicrobial Agents and Chemotherapy, 66(4), e01437-21.

Soumare, H. M., Guelbeogo, W. M., van de Vegte-Bolmer, M., van Gemert, G. J., Soumanaba, Z., Ouedraogo, A., Ouattara, M. S., Abdullahi, A., Jadama, L., Camara, M. M., Gaye, P. M., Mendy, M., Davis, N., Tiono, A. B., D'Alessandro, U., Drakeley, C., Bousema, T., Moreno, M., & Collins, K. A. (2021). Maintaining Plasmodium falciparum gametocyte infectivity during blood collection and transport for mosquito feeding assays in the field. Malaria Journal, 20, 191.

Stanisic, D. I., Liu, X. Q., De, S. L., Batzloff, M. R., Forbes, T., Davis, C. B., Sekuloski, S., Chavchich, M., Chung, W., Trenholme, K., McCarthy, J. S., Li, T., Sim, B. K. L., Hoffman, S. L., & Good, M. F. (2015). Development of cultured Plasmodium falciparum blood-stage malaria cell banks for early phase in vivo clinical trial assessment of anti-malaria drugs and vaccines. Malaria Journal, 14, 143.

Thiam, L. G., Ansah, F., Niang, M., Awandare, G. A., & Aniweh, Y. (2022). Short-term cryopreservation and thawing have minimal effects on Plasmodium falciparum ex vivo invasion profile. Frontiers in Cellular and Infection Microbiology, 12, 997418.

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.

DNA Nanoballs: How MGI's DNBSEQ Rewrites Sequencing

 

Picture a single strand of DNA curling up into a tight little ball, thousands of identical copies of itself packed into a space smaller than a red blood cell. Now imagine spreading billions of these balls across a chip, each one sitting in its own private parking spot, and reading every single one of them at once. That, in essence, is what MGI's DNBSEQ platform does every time it runs. No PCR thermal cycling. No bridge amplification. Just a ball of DNA and a very patient camera.

If you have been following this series, you already know the cast of characters. We started with Sanger sequencing, the original workhorse that gave us the first human genome, one careful read at a time. Then came the next-generation revolution: Illumina's bridge amplification and reversible terminator chemistry, Oxford Nanopore's electrical current reading of DNA threading through a protein pore, PacBio's real-time single-molecule imaging, and Ion Torrent's clever trick of measuring pH instead of light. Today we add a sixth platform to the lineup, one that has quietly become one of the most widely used sequencing technologies outside the United States: MGI's DNBSEQ.

Where did this technology come from?

DNBSEQ traces its roots back to Complete Genomics, a California company that published a landmark paper in Sciencein 2010 describing a method for sequencing a human genome using what they called self-assembling DNA nanoarrays (Drmanac et al., 2010). The idea was radical for its time: instead of amplifying DNA fragments with PCR, which introduces its own errors and biases, why not let the DNA circularize and replicate itself using a gentler, more faithful process? Complete Genomics was later acquired by the Chinese genomics company BGI, and its sequencing chemistry evolved into what we now call DNBSEQ, developed and commercialized by MGI Tech, BGI's technology arm.

Why does it matter that this technology skips PCR? Anyone who has tried to amplify an AT-rich genome like Plasmodium falciparum, sitting at roughly 80 percent AT content, knows the frustration of GC bias, dropout regions, and uneven coverage that standard PCR-based library prep can introduce. A method that avoids repeated PCR cycling has an obvious appeal for exactly this kind of genome.

So how does a DNA nanoball actually form?

The process starts in a familiar place. Double-stranded DNA fragments, each carrying adapter sequences at their ends, are heated until they denature into single strands. From here, DNBSEQ takes a different road than Illumina.

A splint oligonucleotide, a short synthetic sequence complementary to both ends of the same single strand, hybridizes to those ends simultaneously, pulling them together into a nicked circle. DNA ligase then seals the nick, producing a stable, closed single-stranded circle. Think of it like taking a piece of string and tying its two ends together to make a loop.

Once that circle exists, rolling circle amplification takes over. A single high-fidelity polymerase travels around and around the circular template, producing a long concatemer, essentially the same sequence repeated over and over, roughly 100 to 1000 times. This concatemer naturally coils up into a compact, tangled structure: the DNA nanoball.

Why is this gentler than PCR? In PCR, each new copy is made from the previous copy, so any error introduced early in the process gets carried forward and amplified exponentially with every cycle. Rolling circle amplification instead copies the same original circular template again and again. An error introduced during one pass around the circle does not get compounded, because the next pass starts fresh from the same original template. MGI's own technical materials describe this as producing far less accumulated amplification error than PCR-based approaches, and one convenient side effect is that DNB concentration can be measured with a simple Qubit fluorometer, no expensive quantification instrument required.

Getting the nanoballs onto the chip

Once formed, DNBs are loaded onto a patterned array flow cell. Rather than letting DNA molecules land wherever they please on a randomly coated surface, these chips are etched with an orderly grid of binding sites spaced at submicron distances, each designed to hold exactly one DNB. It is a bit like an egg carton for DNA: every nanoball gets its own compartment, evenly spaced from its neighbors. This deliberate spacing helps prevent signal crosstalk between neighboring clusters, which is one of the failure modes that can plague random-cluster sequencing arrays as they get more crowded.

Reading the sequence: cPAS

The actual base calling happens through what MGI calls combinatorial probe-anchor synthesis, or cPAS. Four fluorescently labeled probes, one for each base, bind to the growing strand in a cycle. Lasers excite the fluorophores, a camera captures the resulting flash of color at every single DNB position on the chip, and proprietary software translates millions of these tiny light signals into a base call. The cycle repeats, one base at a time, across the whole flow cell simultaneously.

Where DNBSEQ sits in the platform landscape

MGI now offers a range of instruments spanning very different throughput needs, from compact benchtop systems like the E25 and G99 up through high-throughput workhorses like the G400, T7, and T20×2, the last of which is designed for genome centers running dozens of samples a day.

How does it actually stack up against Illumina, the platform most labs already know? A benchmarking study using the Korean Reference Genome compared DNBSEQ-T7 against six Illumina platforms, including the NovaSeq 6000, and found the two technologies to be broadly comparable for whole-genome sequencing accuracy (Kim et al., 2021). A separate study looking at DNA metabarcoding, sequencing COI gene amplicons from soil samples, found highly correlated results between MGI's DNBSEQ-G400RS and Illumina's NovaSeq 6000, though the MGI platform recovered a somewhat higher number of distinct taxonomic units in that particular comparison (Anslan et al., 2021).

Why this matters for malaria genomics

For a lab working on P. falciparum genomics, sequencing choices are never purely academic. Library prep cost, turnaround time, and how a platform handles an extremely AT-rich genome all factor into which sequencer ends up processing your field isolates. DNBSEQ's PCR-free amplification step is worth understanding even if your lab primarily runs Illumina, because it represents a genuinely different philosophy for how to get from a DNA sample to a finished sequence, and it is increasingly likely to show up in a collaborator's data or a published dataset you are trying to compare against your own.

Related Content

• Sanger Sequencing: The Original Read of Life: https://adwoabiotech.blogspot.com/2026/06/sanger-sequencing-explained-how-one.html

• Illumina Sequencing: Bridges, Clusters, and Light: https://adwoabiotech.blogspot.com/2026/06/what-is-next-generation-sequencing-ngs.html

• Oxford Nanopore: Reading DNA Through a Pore: https://adwoabiotech.blogspot.com/2026/06/what-is-oxford-nanopore-sequencing-how.html

• PacBio vs. Nanopore: Two Roads to Long Reads: https://adwoabiotech.blogspot.com/2026/06/pacbio-smrt-sequencing-vs-oxford.html

• Ion Torrent: When DNA Sequencing Meets pH: https://adwoabiotech.blogspot.com/2026/07/ion-torrent-sequencing-dna-reader-that.html

  References

Anslan, S., Mikryukov, V., Armolaitis, K., Ankuda, J., Lazdina, D., Makovskis, K., Vesterdal, L., Schmidt, I. K., & Tedersoo, L. (2021). Highly comparable metabarcoding results from MGI-Tech and Illumina sequencing platforms. PeerJ, 9, e12254. https://doi.org/10.7717/peerj.12254

Drmanac, R., Sparks, A. B., Callow, M. J., Halpern, A. L., Burns, N. L., Kermani, B. G., Carnevali, P., Nazarenko, I., Nilsen, G. B., Yeung, G., Dahl, F., Fernandez, A., Staker, B., Pant, K. P., Baccash, J., Borcherding, A. P., Brownley, A., Cedeno, R., Chen, L., ... Reid, C. A. (2010). Human genome sequencing using unchained base reads on self-assembling DNA nanoarrays. Science, 327(5961), 78–81. https://doi.org/10.1126/science.1181498

Kim, H.-M., Jeon, S., Chung, O., Jun, J. H., Kim, H.-S., Blazyte, A., Lee, H.-Y., Yu, Y., Cho, Y. S., Bolser, D. M., & Bhak, J. (2021). Comparative analysis of 7 short-read sequencing platforms using the Korean Reference Genome: MGI and Illumina sequencing benchmark for whole-genome sequencing. GigaScience, 10(3), giab014. https://doi.org/10.1093/gigascience/giab014

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.