How Direct RNA Sequencing Works on Oxford Nanopore Pores

Reading RNA the way nature made it

Most RNA sequencing begins with a small act of conversion. You take your RNA, convert it into cDNA with reverse transcriptase, amplify it, and only then do you read it. It works beautifully, and I have used it for years, but every one of those steps leaves fingerprints on the data.

Direct RNA sequencing skips all of that. The native RNA molecule itself goes through the pore. No reverse transcription, no PCR, nothing in between.

The idea is not as new as it might feel. Oxford Nanopore described the approach in a 2016 preprint, and the direct RNA protocol reached customers in April 2017. The full paper appeared in Nature Methods in 2018 (Garalde et al., 2018). Since then the chemistry has moved on, and the current kit is SQK-RNA004, which replaced RNA002.

Three questions come up whenever I explain this to students. How is each pore measured? How many molecules can one pore read? And how much can you get from a whole flow cell?

How is each pore measured individually?

Picture a flow cell as a tiny array of wells. Each well holds a membrane with one protein nanopore embedded in it, and each well has its own electrode wired to a custom chip (an ASIC).

The chip holds a constant voltage across the membrane and measures the ionic current in picoamps, thousands of times per second. For RNA004 the sampling rate is about 4 kHz.

When an RNA molecule carrying a motor-protein adapter is captured, it partly blocks the pore and the current drops to a characteristic level. The motor then ratchets the strand through, 3' end first, at roughly 130 nucleotides per second. As it moves, the current steps up and down depending on which nucleotides (and which modifications) sit in the narrowest part of the pore.

Because every channel has its own electronics, every channel records independently and at the same time. The raw signal is saved per read, tagged with the channel number and read ID, in POD5 format (older runs used FAST5). Basecalling software such as Dorado then converts the signal into sequence.

How many RNA molecules can one pore read?

One at a time. That is the simple answer.

When a strand finishes, it is released, the current returns to the open-pore level, and the pore is free to capture the next molecule. So is there a fixed limit? Not in principle. In practice, the limit is how long the pore stays healthy.

Pores slowly degrade, or get blocked by debris, damaged RNA or secondary structure. The software tries to rescue a blocked pore by briefly reversing the voltage, which sometimes works and sometimes does not. A 1 to 2 kb mRNA takes only about 10 to 15 seconds to read, so a good pore can get through hundreds to a few thousand molecules over a 48 to 72 hour run. Longer RNAs and dirtier samples mean fewer. These are rough figures, and they vary a lot from pore to pore and library to library.

How many molecules can one flow cell read?

That depends on which flow cell you hold in your hand.

• MinION or GridION flow cell: about 2,048 pores sitting on 512 channels. The software only reads 512 at a time. Using a "mux scan" it picks the best pore in each group, then switches to others as the first ones die, so the pores are used up gradually across the run.
• PromethION flow cell: around 3,000 pores across 2,675 channels, with far higher total output.
• Flongle: 126 channels, a small and cheap option that is ideal for testing a library before committing.

For direct RNA on a MinION flow cell, a typical run gives somewhere between one million and several million reads, depending on RNA length, library quality and how many pores stay alive. Treat that as a ballpark, because your own run report is the best guide.

One small note on terminology that trips people up. The kit is the library prep chemistry, and the flow cell is the consumable that carries the pores. They are bought separately, and yield depends on both.

Why bother with all this?

Because the native molecule carries information that cDNA throws away. Direct RNA reads can span a full-length transcript, which helps with isoforms, transcription start sites and alternative polyadenylation. They also let you measure poly(A) tail length on each individual read, and they preserve RNA modifications such as m6A and pseudouridine (Jain et al., 2022; Workman et al., 2019).

The trade-offs are real. You need a fair amount of input (hundreds of nanograms of poly(A)+ RNA), and per-read accuracy is lower than short-read sequencing, although RNA004 narrowed that gap considerably.

Watch the video

Direct RNA sequencing on Oxford Nanopore explained: https://youtu.be/ra1L-pYHXts?si=ymfHktlFAmm-sDzq

Related Content

• PacBio vs Oxford Nanopore: https://adwoabiotech.blogspot.com/2026/06/pacbio-smrt-sequencing-vs-oxford.html

• New to sequencing technologies? Start with Sanger Sequencing Explained: How One of the Oldest DNA Sequencing Methods Still Works to understand the foundational method that long-read platforms eventually built on. 

  If you want to see how short-read NGS stacks up against Sanger, check out What Is Next-Generation Sequencing? NGS vs Sanger Explained. 

  And for a closer look at how Nanopore sequencing compares to the original Sanger method, read What Is Oxford Nanopore Sequencing? How It Differs from Sanger.

References

Garalde, D. R., Snell, E. A., Jachimowicz, D., Sipos, B., Lloyd, J. H., Bruce, M., Pantic, N., Admassu, T., James, P., Warland, A., Jordan, M., Ciccone, J., Serra, S., Keenan, J., Martin, S., McNeill, L., Wallace, E. J., Jayasinghe, L., Wright, C., Blasco, J., Young, S., Brocklebank, D., Juul, S., Clarke, J., Heron, A. J., & Turner, D. J. (2018). Highly parallel direct RNA sequencing on an array of nanopores. Nature Methods, 15(3), 201–206.

Jain, M., Abu-Shumays, R., Olsen, H. E., & Akeson, M. (2022). Advances in nanopore direct RNA sequencing. Nature Methods, 19(10), 1160–1164.

Workman, R. E., Tang, A. D., Tang, P. S., Jain, M., Tyson, J. R., Razaghi, R., Zuzarte, P. C., Gilpatrick, T., Payne, A., Quick, J., Sadowski, N., Holmes, N., de Jesus, J. G., Jones, K. L., Soulette, C. M., Snutch, T. P., Loman, N., Paten, B., Loose, M., Simpson, J. T., Olsen, H. E., Brooks, A. N., Akeson, M., & Timp, W. (2019). Nanopore native RNA sequencing of a human poly(A) transcriptome. Nature Methods, 16(12), 1297–1305.

How to Thaw Plasmodium falciparum Cultures: Step by Step

You pull a vial out of liquid nitrogen, thaw it, wash it, set up the flask, and then you wait. Day one, the smear is empty. Day two, still nothing. Did you just waste a precious stabilate? Almost certainly not, but the anxiety is real, and it is the reason this post exists.

Thawing Plasmodium falciparum is not difficult. It is, however, unforgiving in a few very specific places, and knowing where those places are makes all the difference between a culture that bounces back and one that quietly disappears. Today I will walk you through the full procedure for a stabilate frozen in glycerolyte, explaining the reason behind each step as we go.

A short detour: why glycerol, and why so much care?

The cryoprotective power of glycerol was discovered by accident in 1949, when a bottle on a lab shelf turned out to hold glycerol rather than the albumin its label promised, and the sperm samples stored in it survived freezing. Parasitologists later borrowed the idea, and glycerol-based freezing solutions became the standard way to bank malaria parasites.

Here is the catch. Glycerol protects red cells from ice damage by sitting inside them at high concentration. When you thaw the vial, that glycerol is still in there, so if you suddenly dilute the cells in plain medium, water rushes in faster than the glycerol can leave and the cells burst. The whole thaw is designed around one idea: take the glycerol out gently, by stepping the salt concentration down in stages.

What you need before you start

Warm everything to 37°C (or at least room temperature) and have your fresh O+ red cells washed and ready before the vial leaves the freezer. Scrambling for reagents mid-thaw is how mistakes happen.

Reagent	Volume (per 1 mL of stabilate)	What it does
12% NaCl	0.2 V (200 µL)	First osmotic step, draws water out of the cells before glycerol leaves
1.8% NaCl	10 V (10 mL)	Second step, lets the cells relax towards normal tonicity
0.9% NaCl / 0.2% glucose (optional)	10 V (10 mL)	Gentler final step, useful for old or low-parasitaemia stabilates
Incomplete medium (RPMI 1640)	10 V (10 mL) per wash	Removes the remaining salt
Complete medium	2 to 3 mL	Culture start, with Albumax or serum

V is simply the volume of your thawed stabilate. A typical vial holds about 1 mL, so 0.2 V is 200 µL, but scale everything to your own vial.

Step by step

Step 1: Thaw quickly. Take the vial from liquid nitrogen (or -80°C) and warm it in a 37°C water bath or in your hand until it has just thawed. Wipe the vial with ethanol and move it into the biosafety cabinet. Slow thawing lets ice crystals regrow and shred the cells, so speed matters here.

Step 2: Transfer and measure. Pipette the stabilate into a sterile 15 mL tube and note the volume. That number is your V.

Step 3: Add 12% NaCl drop by drop. Add 0.2 V (about 200 µL for a 1 mL vial) one drop at a time, flicking or swirling the tube gently between drops. Then wait 5 minutes. If I could put a flashing warning on any step, it would be this one. The cells are at their most fragile right now, and dumping the salt in quickly is the single most common cause of a failed thaw.

Step 4: Add 1.8% NaCl slowly. Add 10 V (about 10 mL), dropwise at first and then a little faster as the tube fills, mixing as you go. Spin at 500 × g for 5 minutes.

Step 5: Remove the supernatant. Aspirate carefully and leave a little liquid above the pellet, because the pellet will be small and easy to suck up by mistake. If your stabilate is old, or you suspect the starting parasitaemia was low, add 10 mL of 0.9% NaCl / 0.2% glucose here, mix gently, and spin again at 500 × g for 5 minutes.

Step 6: Wash. Resuspend the pellet in 10 mL of incomplete medium, spin at 500 × g for 5 minutes, and remove the supernatant. Repeat once more if you like. Two gentle washes are plenty.

Step 7: Start the culture. Resuspend the pellet in 2 to 3 mL of complete medium and add fresh red cells to reach about 3 to 5% haematocrit. This step is not optional. A thawed stabilate contains very little blood, and without fresh cells the parasites that survive have nowhere to invade. Keeping the volume small also keeps parasites and red cells close together, which helps that first invasion.

Step 8: Gas and incubate. Gas the flask with 5% CO₂, 5% O₂, 90% N₂ (or use a candle jar if that is what your lab has) and incubate at 37°C.

The waiting game

Change the medium every day, gently, without stirring up the cells that have settled at the bottom. Make a Giemsa smear at 24 hours and again at 48 hours, but do not panic if both are empty. Recovery is rarely dramatic. Parasites often reappear only after 3 to 5 days, once the first round of reinvasion has finished and the numbers are high enough to find under the microscope.

Once you can see parasites, keep the culture at 2 to 5% haematocrit and subculture with fresh red cells when parasitaemia reaches about 1 to 3%. Resist the urge to synchronise or otherwise stress the culture until it is growing steadily. A culture that has just come back from the freezer deserves a quiet few weeks.

When things go wrong

Nothing after 7 to 10 days? The stabilate may have been of low quality or frozen at a very low parasitaemia. Before giving up, keep the culture going and add fresh red cells every week. Some stabilates are simply slow.

Lots of debris or haemolysis in the first pellet? The salt went in too fast, or the vial thawed too slowly. Next time, slow down the 12% NaCl step and use fewer, smaller drops.

Parasites visible but not expanding? Look at your Albumax or serum lot and check the pH of your medium. Growth problems after a successful thaw are very often medium problems in disguise.

Contamination? Discard it promptly and thaw a second vial if you have one.

The short version

Thaw fast, add salt slowly, wash gently, give the parasites fresh red cells, and be patient. If you remember only one thing, remember the 12% NaCl step. It is the moment the whole procedure turns on.

Related Content

• How to Isolate Red Blood Cells for Plasmodium Culture
• The Plasmodium falciparum Life Cycle Explained
• How to Calculate Hematocrit for Passaging
• Giemsa Stain: Morphology and History
• DNA Sequencing Platforms Compared: Sanger, Illumina, Nanopore, PacBio

Plasmodium falciparum WGS: How to Prepare Red Blood Cells

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.

Related Content

• How to Isolate Red Blood Cells for Plasmodium Culture
• The Plasmodium falciparum Life Cycle Explained
• How to Calculate Hematocrit for Passaging
• Giemsa Stain: Morphology and History
• DNA Sequencing Platforms Compared: Sanger, Illumina, Nanopore, PacBio

ANOVA in Excel for Beginners: Mean, SD and Post-Hoc Tests

Meeting someone new follows a script. How are you? What do you do? Where did you study? Nobody skips it, because those small questions build the ground for anything more interesting. Data, I have come to think, deserves exactly the same courtesy.

Before you throw a statistical test at your results, you greet them. What is your average? How scattered are you? What do you look like on a graph? Only then do you ask the big question: are you really different from each other? This post walks through that conversation in Excel, using one control and three experimental conditions.

The walkthrough is adapted from the Adwoa Biotech YouTube tutorial. I have tightened a few statistical points along the way.

Step 1: Meet the average

Say you have seven readings for each of four groups: a control and three conditions. The mean gives every group one representative number. You add up all the values, divide by how many there are, and in Excel that is simply =AVERAGE(...).

One warning, though. When your values are wildly scattered, the mean becomes a poor spokesperson. For skewed data, the median (reported with the interquartile range) tells a more honest story.

Step 2: Ask about the spread

An average on its own is a sentence without context. Two groups can share the same mean and behave completely differently, so you also ask how far the readings wander from it. That is the standard deviation, and for a sample you want =STDEV.S(...).

You will often see the standard error of the mean (SEM) too, which is the SD divided by the square root of n. They answer different questions, so it helps to keep them apart:

• SD describes how scattered your data are.
• SEM describes how precisely you have pinned down the mean.

Whichever one you plot, say so in the figure legend. Your readers should never have to guess.

Step 3: Draw it

Our four groups are independent of one another, so a bar chart fits well. Had they been time points, I would reach for a line graph to show the progression instead.

In Excel, go to Insert and pick a bar chart. Then comes the part I enjoy most: decluttering. The rule is simple, and a little ruthless. If an element does not help communicate the data, it goes. Gridlines leave first, and the cleaner chart feels a bit like opening a window in a stuffy room.

After that, add what a stranger needs in order to read it:

• Axis titles that name the quantity and its units, for example "Absorbance (405 nm)", with "Experimental condition" on the x-axis.
• A title that tells the viewer what the chart is about.
• Error bars, with one catch. Excel's built-in "Standard Deviation" option calculates across the plotted bars rather than within each group, so the result is meaningless here. Choose Custom and point it at the SD (or SEM) cells you calculated yourself.

With small samples like n = 7, consider overlaying the individual data points as well. Bars can hide a surprising amount of drama.

Step 4: Are they really different?

For two groups, a t-test does the job, and in Excel the Welch version (unequal variance) is the safer default. With four groups, though, it is tempting to run a t-test on every pair. Resist that temptation.

Why? Four groups give six pairwise comparisons, and each test is another roll of the dice for a false positive. At α = 0.05 across six tests, the chance of at least one fluke is 1 − 0.95⁶, which is roughly 26%. That inflated error rate is exactly why we use analysis of variance (ANOVA): one test across all the groups, with the false-positive rate held at 5%.

Switching on the Analysis ToolPak

Excel keeps this tool behind a door, and nearly everyone looks for it in the wrong place at least once.

1. Go to File, then Options, then Add-ins.
2. At the bottom, set Manage to Excel Add-ins and click Go.
3. Tick Analysis ToolPak.

A Data Analysis button now appears on the Data tab. It lives under Add-ins in the options menu, not under Data, which is where the trap is.

Feeding it the right food

ANOVA does not want your means and SDs. It needs the raw values for every replicate, because the variance is precisely what the test is measuring. If your numbers come from a standard curve, convert each replicate to a concentration individually before you start.

• Put each group in its own column, with a heading.
• Paste values, not formulas, when moving data between cells.
• If you include headings, tick Labels in first row, otherwise Excel complains about non-numeric data.
• Choose an output range yourself, because Excel will not pick one for you.

Then select Anova: Single Factor. Alpha is set to 0.05 by default, which is the usual convention. A stricter threshold would be 0.01.

Reading the output

You get a summary table (count, sum, average and variance for each group) and an ANOVA table. The second one is the part that makes people nervous, so here is what each column means:

Term	What it means	In our example
SS	Sum of squares: total variability, split into between and within groups	Reported for each
df	Degrees of freedom. Between = groups − 1. Within = observations − groups	3 and 24 (28 − 4)
MS	Mean square: SS divided by df, an average amount of variation	Reported for each
F	MS between divided by MS within	One value
P-value	Probability of data this extreme if the null hypothesis were true	About 0.0019

Notice that there is only one p-value. That is because there is only one test. The "within groups" row is not a second result, it is simply the yardstick the F ratio is measured against.

What a p-value does and does not say

Our p of 0.0019 sits below 0.05, so we reject the null hypothesis that all the group means are equal. Two misreadings are very common, and both are worth avoiding:

• Alpha is not confidence in the null hypothesis. Setting α = 0.01 does not mean you are 99% sure nothing is happening. It means you accept a 1% false-positive rate when there truly is no effect.
• A p-value is not "the probability it was all chance." It is the probability of seeing data at least this extreme if the null hypothesis were true.

The catch: ANOVA does not say who

This is the part that gets skipped most often. ANOVA is an omnibus test. It tells you that at least one group differs, and then it shrugs. Did condition 1 beat the control? Are conditions 2 and 3 the odd pair? The test cannot say.

To find out, you need a post-hoc test:

• Tukey's HSD compares every pair of groups.
• Dunnett's test compares each condition against the control, which is often exactly what an experiment needs.

The ToolPak includes neither, so you will need other software such as R, GraphPad Prism or JASP.

Before you trust it

ANOVA assumes the observations are independent, the residuals are roughly normal, and the group variances are similar. With seven values per group, those assumptions are hard to verify. If they look shaky, the Kruskal-Wallis test is the non-parametric alternative, although Excel does not have it built in either.

The short version

1. Say hello: mean, SD (or median and IQR), and a clean chart.
2. Run ANOVA on the raw replicates, never on the summaries.
3. Read the p-value carefully.
4. Follow up with a post-hoc test to learn which groups differ.
5. Tell your readers whether the error bars show SD or SEM.

Video callout: Want to see the clicks in action? Watch the original walkthrough here: https://www.youtube.com/watch?v=3jYo1W6hyN0

Related Content

• Calculating log2 and log10 values

5% Haematocrit in P. falciparum Culture: Volumes and Why

When I was culturing parasites in the field, one number followed me everywhere: 5%. Five percent haematocrit. It sounds like a small, fussy detail, doesn’t it? Yet it quietly decides whether a culture thrives or collapses.

What haematocrit actually means

Haematocrit is simply the proportion of a suspension that is packed red blood cells. In a culture at 5% haematocrit, red cells make up a twentieth of the total volume, and everything else is culture medium. Think of it as a very thin cellular soup, mostly broth with a modest scattering of cells.

Why not more cells?

Plasmodium falciparum lives inside red blood cells, so you might assume that more cells means more parasites. In practice, a denser culture burns through nutrients quickly, acidifies the medium, and leaves you changing it far more often than you would like. In the field, where supplies are limited and the power can be unpredictable, that matters a great deal.

Why not fewer?

Too sparse, and merozoites struggle to find fresh cells to invade after the schizont bursts. Growth slows, and the parasitaemia you worked so hard to establish stalls. Five percent is a compromise: enough cells for efficient reinvasion, enough medium to keep conditions stable between changes.

What 5% looks like at the bench

Here are two culture sizes I used regularly. In both, the volume is the total culture volume, so the cells are part of it, not added on top.

• For a 5 mL culture: 250 µL of packed red cells (infected cells included) with 4.75 mL of complete medium.
• For a 10 mL culture: 500 µL of packed red cells (infected cells included) with 9.5 mL of complete medium.

If you work from a 50% haematocrit stock instead, double the cell volume: 500 µL of stock for a 5 mL culture, or 1 mL for a 10 mL culture. Top up with complete medium to the final volume.

A few habits that helped me

Always start from properly washed, packed cells, after a clean spin with the supernatant and buffy coat removed. Packed cells are viscous, so use a wide-bore or cut tip and pipette slowly. If you find small volumes of packed cells awkward to measure, make a more dilute stock first and work from that. It is easier to be accurate with a larger volume, and consistency from one passage to the next is what keeps a culture healthy.

The takeaway

Five percent is not magic, but it is a reliable starting point that balances parasite growth against nutrient supply. Once you understand why it works, you can adjust it sensibly when conditions call for something different.

Video callout: Want to see this in practice? Watch the walkthrough on the Adwoa Biotech YouTube channel: https://youtu.be/AF1bkbbgf9w?si=_9ICA7np-ECNXp4s

Related Content

• Hematocrit calculation for passaging
• RBC isolation protocol
• P. falciparum life cycle

MagMAX RNA Isolation Protocol for Tempus Blood RNA Tubes

 

MagMAX for Stabilized Blood Tubes RNA Isolation Kit Protocol for Tempus Tubes (Cat. 4451893)

This is the manufacturer's protocol for purifying total RNA, including microRNA, from human whole blood collected in Tempus Blood RNA Tubes, paraphrased from the Applied Biosystems protocol (Part No. 4452007, Rev. A). It works with magnetic beads instead of spin columns, and you can run it in 1.5 mL tubes or in the 96-well Processing Plate supplied with the kit. Expect roughly 3 to 25 µg of RNA from 3 mL of blood.

Before you start

• Draw 3 mL of blood into the Tempus tube and shake or vortex it for 10 seconds straight away. Poor mixing means poor stabilization and microclots.
• Stabilized blood keeps for up to 5 days at 18 to 25 °C, up to 7 days at 4 °C, and long term at −20 to −80 °C (keep it off direct contact with dry ice).
• Prepare Wash Solution 1 by adding 8 mL of 100% isopropanol to each bottle of concentrate. Prepare Wash Solution 2 by adding 24 mL of 100% ethanol to the concentrate. Invert 3 to 4 times, mark the bottles, and store at room temperature. Each isolation uses 300 µL of each.
• You need a refrigerated swing-bucket centrifuge with 50 mL tube adapters that can reach 5,000 × g at 4 °C, a magnetic stand, and either a vortexer with a 1.5 mL tube adapter or an orbital plate shaker.
• Vortex adapter settings used by the manufacturer (scale of 1 to 10): 4 to 5 for binding and washes, 6 to 7 for digestion and elution. Orbital plate shaker settings: 6 to 7 for binding and washes, 8 to 9 for digestion, drying and elution. Test your own equipment with water so nothing spills.
• Wear gloves, use RNase-free tips, and wipe the bench and pipettes with an RNase decontamination solution.

Dilute the stabilized blood

1. If the sample is frozen, thaw the Tempus tube on ice for 10 to 30 minutes. Keep samples on ice as much as possible.
2. Pour the whole contents into a labeled 50 mL conical tube.
3. Add 3 mL of Tempus 1× PBS to reach 12 mL. If you collected less than 3 mL of blood, top up with PBS to 12 mL, otherwise yields drop sharply.
4. Cap tightly and vortex at vigorous speed for 30 seconds. Froth is normal. Go straight to the next section.

Pellet and wash the crude RNA

1. Centrifuge at 5,000 × g for 15 minutes at 4 °C. The pellet is transparent and invisible, so handle the tubes gently.
2. Pour off the supernatant.
3. Mix the Tempus Pre-Digestion Wash by gently inverting the bottle 3 to 4 times. Add 4 mLto each tube, recap, and vortex at moderate speed for 5 seconds, enough to draw the liquid about three-quarters of the way up the tube.
4. Centrifuge again at 5,000 × g for 10 minutes at 4 °C.
5. Decant the supernatant carefully. Decanting is recommended because a pipette can take the pellet with it.
6. Drain the tubes inverted on absorbent paper for 2 minutes, then blot the rims.
7. Keep the tubes on ice and move on immediately.

Resuspend the pellet and digest with protease and TURBO DNase

Just before use, prepare the resuspension mixture. Scale it by the number of samples and add 5% extra to cover pipetting loss.

Component	Volume for 1 sample
Tempus Resuspension Solution	117.5 µL
Tempus Proteinase	2.5 µL
Total	120 µL

1. Mix the components by vortexing at moderate speed, spin briefly for 2 to 3 seconds, and keep on ice.
2. Add 120 µL of the mixture to each conical tube, placing it at the bottom of the tube, and recap.
3. Vortex at gentle speed for 5 to 10 seconds to resuspend the pellet.
4. Transfer the full volume to a 1.5 mL microfuge tube or to one well of the 96-well Processing Plate. You can pause here and hold the resuspended pellets on ice for up to 4 hours (cover the plate wells).
5. Add 10 µL of TURBO DNase to each tube or well and mix at vigorous speed for 10 minutes. Do this at 18 to 25 °C. If your lab is colder than 18 °C, use a 25 °C incubator.

Bind the RNA to the beads

1. Vortex the RNA Binding Beads at vigorous speed until fully resuspended.
2. To each tube or well, add 50 µL of Binding Solution Concentrate and 20 µL of the bead suspension. Add the beads before the isopropanol.
3. Mix at moderate speed for 1 minute.
4. Add 200 µL of 100% isopropanol and mix at moderate speed for 3 minutes.
5. Capture the beads on the magnetic stand for about 1 to 3 minutes, until the solution clears.
6. Aspirate and discard all the supernatant without disturbing the beads.
7. Take the tubes or plate off the magnet. This is critical before the next step.

Wash twice with Wash Solution 1

1. Add 150 µL of Wash Solution 1 and mix at moderate speed for 1 minute.
2. Capture the beads for about 1 to 2 minutes and discard the supernatant.
3. Remove from the magnet, then repeat the wash once.

Wash twice with Wash Solution 2

1. Add 150 µL of Wash Solution 2 and mix at moderate speed for 1 minute.
2. Capture the beads for about 1 to 2 minutes and discard the supernatant.
3. Remove from the magnet, then repeat the wash once.

Dry the beads

• Tubes: open them and invert on absorbent paper for 2 minutes at room temperature. If liquid remains, remove it with a fine-tipped pipette without touching the beads, then invert for 1 minute more.
• Plate: shake at vigorous speed for 2 minutes at room temperature. If liquid remains, shake for 2 minutes more.
• Do not over-dry the beads, and make sure no alcohol is left before elution.

Elute the RNA

1. Add 80 µL of Elution Buffer (you can use anywhere from 20 to 80 µL depending on the concentration you want) and mix at vigorous speed for 4 minutes.
2. Capture the beads for about 1 to 3 minutes.
3. Transfer the supernatant to a nuclease-free tube without disturbing the beads. Keep it, because the purified RNA is in this supernatant.
4. Store on ice for immediate use, at −20 °C for up to 6 months, or at −80 °C for the long term.

Checking yield and quality

• Expect an A260/A280 ratio of 1.8 to 2.2. Values outside that range can still work for qRT-PCR.
• For human blood RNA from this kit, a 28S to 18S rRNA ratio of 1.0 or higher is considered acceptable. A RIN from a bioanalyzer is the more reliable measure of integrity.

Troubleshooting

• Low or no yield: check that the tube was filled to the black mark, that the blood was diluted to a final 12 mL with PBS and vortexed for 30 seconds, and that you used the full centrifuge times and speeds in a 50 mL tube in a swing-bucket rotor. Lysate that was exposed to more than 37 °C can be frozen, thawed and repurified. Also check that the beads were fully resuspended, were never frozen, and were added before the isopropanol.
• Lost pellet or beads: decant slowly rather than pipetting off the pellet, and aspirate slowly with the tip away from the captured beads.
• Beads clump during elution: warm the plate or tubes at 70 °C for 5 minutes and repeat the 4-minute mix. Next time, preheat the Elution Buffer to 70 to 80 °C. If the beads were over-dried, extend the elution mixing to 10 minutes.
• Bead carryover (light brown eluate): capture again on the magnet and move the RNA to a fresh tube. A small amount does not inhibit RT or RT-PCR.
• Too much DNA: make sure the digestion ran for the full 10 minutes at vigorous mixing and at 18 to 25 °C, and that the pellet was washed and drained well so stabilizing reagent was not carried over.
• RNA fails downstream: salt or alcohol carryover is the usual cause, so check that the correct alcohol and volume went into each wash concentrate, remove every supernatant fully, and let the beads dry before elution. A green or reddish eluate signals inhibitors, so spin it at 16,000 × g for 3 minutes and move the clear fraction to a new tube.

Source: Applied Biosystems, MagMAX for Stabilized Blood Tubes RNA Isolation Kit (Compatible with Tempus Blood RNA Tubes) Protocol, Part No. 4452007, Rev. A (2010).