SYBR Green I Assay for Malaria Drug Testing: Step by Step

Here is a fact I never get tired of: the cell that malaria lives in has thrown away its own DNA. Mature human red blood cells have no nucleus and no genome, so there is nothing in them to stain. That means any DNA you find in a Plasmodium falciparum culture belongs to the parasite. And what does that allow us to do? If we can make DNA glow, we can count parasites, and if we can count parasites, we can ask how much of a drug it takes to stop them.

For a long time, answering that question meant working with radioactivity. Labs fed parasites a radiolabelled substrate and measured how much they took up, which demanded special equipment, safety procedures and a lot of patience. Then, in 2004, Smilkstein and colleagues reported a simple, inexpensive, one-step alternative built on SYBR Green I, a dye whose fluorescence rises sharply when it binds double-stranded DNA (Smilkstein et al., 2004). No isotopes, no wash steps. Just a plate reader. 

In this post I walk through the whole assay, from the drug dilution plate to the IC50, using the layout from my video. It is a 96-well format, so it scales nicely from a single compound to a full screen.

What the assay actually measures

SYBR Green I barely fluoresces on its own. Slip it between the strands of DNA and it lights up, roughly a thousand (1000) times brighter. After 72 hours of growth, you lyse the cells, add the dye, and read the plate. More parasites mean more DNA, which means more green. Add an effective drug and the signal falls. Plot fluorescence against the logarithm of the drug concentration, fit a curve, and the midpoint is your IC50.

One small note on timing. The original paper measured growth after 48 hours, while many labs now use 72 hours so that the parasites go through more than one cycle and slower-acting drugs have a fair chance to show an effect. Pick one, and keep it identical across experiments.

Step 1: Build the drug plate

Prepare your drugs at twice the final concentration in complete medium, because they will be diluted one to one when the parasites go in. Dissolve compounds in DMSO if needed, and keep the final DMSO at or below 0.2% in every well, including the controls.

Add 50 µL of complete medium to the wells that will carry the lower concentrations. Put 100 µL of the top 2× concentration in the first drug column, then transfer 50 µL to the next column, mix, and carry on across the plate, discarding the last 50 µL. Every drug well now holds 50 µL, and each column is half the concentration of the one before. That is your two-fold serial dilution.

Then plan the controls. Edge wells evaporate faster and give odd readings, so fill them with 100 µL of water or PBS and do not use them for parasites. In my layout, columns 2 to 10 of rows B to D hold the test compound, rows E to G hold chloroquine as a reference drug, and column 11 holds the controls: no-drug parasites for 100% growth, and uninfected red cells for background.

Step 2: Prepare the parasite suspension

Hematocrit is the percentage of the culture volume that is packed red cells. We want 2% in the well. Because the parasites are mixed one to one with the drug, you prepare the suspension at 4% hematocrit and the well ends up at 2% in a total volume of 100 µL.

Do you need to add uninfected blood? Only if your culture is above the parasitaemia you want. Parasitaemia is the fraction of red cells that are infected, so the infected cells are already part of the hematocrit and are not counted twice. If your culture is already at 0.5% to 1%, spin it at 500 × g for 5 minutes, remove the supernatant, and use the pellet as your packed cells. If it is higher, dilute the pellet with uninfected red cells until you reach the target parasitaemia.

Here is a bulk mix for one plate, with a little extra for pipetting losses:

Component	Volume (5.5 mL batch at 4% hematocrit)
Packed infected red cells (pellet)	220 µL
Complete medium	5.28 mL

Make a small separate batch of uninfected red cells at 4% hematocrit for the background wells, from the same donor and batch you used to dilute the parasites. That way the background matches the cells in the rest of the plate.

Step 3: Combine and incubate

Add 50 µL of the parasite suspension to each drug well and to the no-drug wells, and 50 µL of the uninfected suspension to the background wells. Each well now holds 100 µL at 2% hematocrit. Place the plate in a humid chamber, flush it with a low-oxygen gas mixture (5% O₂, 5% CO₂, 90% N₂), seal it, and incubate at 37 °C for 72 hours. This is a good time to go and do something else. The plate does not need you.

Step 4: Lyse and stain

The lysis buffer does two jobs. Saponin and Triton X-100 open the red cells and the parasite membranes, and EDTA keeps nucleases quiet, so that the dye can reach the DNA. Prepare it in advance, and add the dye only on the day.

Component	Final concentration
Tris, pH 7.5	20 mM
EDTA	5 mM
Saponin	0.008% (w/v)
Triton X-100	0.08% (v/v)
SYBR Green I (10,000× stock)	0.2 µL per mL of lysis buffer (about 2×)

This is the recipe from the original paper, and it works well for most labs (Smilkstein et al., 2004). SYBR Green I is light sensitive, so protect the buffer with foil. Add 100 µL to every well and mix until no visible red cell sediment remains. Then wrap the plate in foil and leave it for 1 hour at room temperature in the dark.

Step 5: Read the plate

Use a fluorescence plate reader with excitation at about 485 nm and emission at about 530 nm. Black plates reduce crosstalk between wells. After three days of waiting, the reading takes about a minute, which feels almost rude.

Step 6: From fluorescence to IC50

First, subtract the mean of the uninfected red cell wells from every reading. Second, set the mean of the no-drug wells to 100% growth. Third, plot the percentage growth against the log of the drug concentration and fit a sigmoidal dose-response curve with nonlinear regression, in GraphPad Prism or a similar program. The concentration at the midpoint of the curve is your IC50.

Run each concentration in triplicate, and repeat the whole experiment on at least two or three separate days. Technical replicates tell you how well you pipette, while independent experiments tell you whether the result is real. And check your reference drug: if the chloroquine IC50 for a well-characterised strain is far from the values reported in the literature, the assay is telling you something, and it is not about your compound.

Things that bite you

• White blood cells. They carry DNA too. This is no problem for cultured parasites, but it matters for fresh patient blood unless you remove the leukocytes first.
• DNA-binding compounds. A drug that grabs DNA competes with the dye and can fake a result, and a coloured compound can quench the signal.
• Slow-acting drugs. Some compounds need a longer exposure than 72 hours to show their full effect.
• Edge wells. Fill them with water or PBS, not parasites.
• Light and age. Protect the dye from light and make the final buffer fresh.

Watch the video

Prefer to see it done? I walk through the whole assay, step by step, in the video below, including the plate layout and the dose-response curve.

Video: SYBR Green I Assay: Malaria Drug Testing Step by Step

Related Content

• Hematocrit calculation for passaging P. falciparum
• The P. falciparum life cycle
• Giemsa stain: morphology and history
• Red blood cell isolation protocol

References

Smilkstein, M., Sriwilaijaroen, N., Kelly, J. X., Wilairat, P., & Riscoe, M. (2004). Simple and inexpensive fluorescence-based technique for high-throughput antimalarial drug screening. Antimicrobial Agents and Chemotherapy, 48(5), 1803-1806. https://doi.org/10.1128/AAC.48.5.1803-1806.2004