A few weeks ago I put together a video on how to isolate and wash red blood cells for Plasmodium falciparum culture. It covers plasma removal, buffy coat removal, three washes, and a tidy 5% hematocrit at the end. Then a question landed that I had not planned for: what changes if the culture is going to be used for whole genome sequencing (WGS)?
The honest answer is: quite a lot. I expected a tweak or two. What I got was a rethink of nearly every stage, from the donor blood right through to the way the parasites are harvested. So let's walk through it together.
Why does sequencing change the rules?
For a normal culture, a few stray white blood cells are a nuisance at worst. They do not stop the parasites from growing, and nobody is counting them. Sequencing is a different story, because every cell in the flask contributes DNA to the tube, and not all DNA is equal.
Think about the numbers. The P. falciparum genome is only about 23 million base pairs. A single human white blood cell carries roughly 6 billion. That means one leukocyte holds around 250 times more DNA than a haploid ring-stage parasite. A handful of survivors from the buffy coat can quietly take over your sequencing reads, and you only find out after you have paid for the run. Painful.
Step 1: Remove white blood cells properly
In the standard video protocol we aspirate the buffy coat and move on. For WGS, that is not enough. Pass the blood through a leukocyte-depletion filter such as a Plasmodipur filter, or through a cellulose column such as CF11 (Auburn et al., 2011; Venkatesan et al., 2012). This is the single biggest change on the list.
But there is another part that is easy to miss. It applies to the uninfected donor red blood cells you add to the culture, not only to patient blood. If the donor cells carry white blood cells, those white cells come along for the ride into every passage.
Step 2: Be choosy about the donor blood
Use fresh, well-characterised donor blood, ideally group O positive, and aim to use it within about 2 weeks of collection. Keep the washing steps from the original protocol: three washes in incomplete RPMI 1640. Where you can, use the same red blood cell batch across related cultures, so that batch-to-batch variation does not muddy the comparison.
Step 3: Keep the culture short
The longer parasites sit in a flask, the more they adapt to it. Culture adaptation can select for particular clones, and it has been linked to loss-of-function mutations and changes in subtelomeric and var gene regions (Claessens et al., 2017). If you are sequencing fresh patient isolates and want to preserve the original diversity, keep it to a few cycles.
Then ask yourself a simple question: do you want one genotype or many? Limiting dilution gives you a single clone, which makes variant calling much cleaner. A mixed infection gives you several clones in one tube, which is biologically interesting but harder to untangle.
Two housekeeping points while you are here:
- Test the culture for mycoplasma. Contamination adds foreign DNA that you do not want in the library.
- If you need a defined stage, synchronise with 5% sorbitol for about 10 minutes at 37 °C. Rings carry one genome copy, while late schizonts carry many and give you more DNA per cell.
Step 4: Harvest with saponin, not just a pellet
In the culture-only workflow, you simply keep the cells growing. For DNA, you need to get rid of the red blood cell contents first. Lyse the red blood cells with saponin at about 0.05% to 0.15% for roughly 5 minutes. This frees the parasites while the haemoglobin and most host material stay in solution.
Next, spin the parasites down (typically 2,000 to 3,000 g for 5 to 10 minutes) and wash several times in cold PBS until the supernatant runs clear. Adjust the speeds to your own centrifuge and tubes. For the best yield, harvest at high parasitaemia, around 5% or more, when the parasites are at late trophozoite or schizont stage.
Step 5: What if the DNA yield is low?
Sometimes you simply cannot grow enough parasites, especially with field isolates at low parasitaemia. There are two main rescue routes:
- Selective whole genome amplification (sWGA) preferentially amplifies parasite DNA over human DNA (Oyola et al., 2016). It is very helpful for low-input samples, but it can introduce coverage bias, so it is not ideal if copy number or structural variants are your focus.
- CpG-methylation-based host depletion, for example with the NEBNext Microbiome DNA Enrichment kit, takes advantage of a neat fact: human DNA is CpG-methylated, while P. falciparum DNA is not.
Step 6: Check before you commit
Quantify your DNA with a fluorometric method such as Qubit, not NanoDrop, because NanoDrop overestimates in the presence of contaminants. Then check the human-to-parasite ratio with qPCR using a human target and a Pf target. A small pilot run before the full sequencing batch can save a lot of money and heartache.
Culture versus WGS at a glance
| Stage | Routine culture | Culture for WGS |
|---|---|---|
| White cell removal | Remove buffy coat | Buffy coat removal plus Plasmodipur or CF11 filtration |
| Donor blood | Washed three times | Fresh, filtered, same batch where possible |
| Culture length | As long as needed | As short as possible |
| Harvest | Keep cells growing | Saponin lysis, cold PBS washes |
| Quality control | Giemsa smear | Qubit plus human-to-parasite qPCR, mycoplasma test |
Watch the video
If you would like to see the standard red blood cell isolation workflow first, the full video is here: Isolating Red Blood Cells for Plasmodium Culture. It is the foundation for everything above, and the WGS changes slot neatly on top of it.
The take-home message
Culture asks, "Are my parasites happy?" Sequencing asks, "Whose DNA is in this tube?" Once you think about it that way, the changes make sense: filter the white cells, keep the culture short, lyse with saponin, and check the human-to-parasite ratio before you spend a cent on sequencing.
References
Auburn, S., Campino, S., Clark, T. G., Djimde, A. A., Zongo, I., Pinches, R., Manske, M., Mangano, V., Alcock, D., Anastasi, E., Maslen, G., Macinnis, B., Rockett, K., Modiano, D., Newbold, C. I., Doumbo, O. K., Ouédraogo, J.-B., & Kwiatkowski, D. P. (2011). An effective method to purify Plasmodium falciparum DNA directly from clinical blood samples for whole genome high-throughput sequencing. PLoS ONE, 6(7), e22213.
Claessens, A., Affara, M., Assefa, S. A., Kwiatkowski, D. P., & Conway, D. J. (2017). Culture adaptation of malaria parasites selects for convergent loss-of-function mutants. Scientific Reports, 7, 41303.
Oyola, S. O., Ariani, C. V., Hamilton, W. L., Kekre, M., Amenga-Etego, L. N., Ghansah, A., Rutledge, G. G., Redmond, S., Manske, M., Jyothi, D., Jacob, C. G., Otto, T. D., Rockett, K., Newbold, C. I., Berriman, M., & Kwiatkowski, D. P. (2016). Whole genome sequencing of Plasmodium falciparumfrom dried blood spots using selective whole genome amplification. Malaria Journal, 15, 597.
Venkatesan, M., Amaratunga, C., Campino, S., Auburn, S., Koch, O., Lim, P., Uk, S., Socheat, D., Kwiatkowski, D. P., Fairhurst, R. M., & Plowe, C. V. (2012). Using CF11 cellulose columns to inexpensively and effectively remove human DNA from Plasmodium falciparum-infected whole blood samples. Malaria Journal, 11, 41.
