Dilution Series

You need to set up a dilution series whenever you want to quantify bacteria or phages on a plate. If there are too many colonies or plaques on a plate, you can't count them anymore (or it takes forever), so you need to dilute your sample beforehand and at the end factor the dilution into your calculation of the concentration your sample has.

Setting up a dilution series

In most cases you want to use a 96-well microplate for your dilution series. Together with a multi-channel pipette, you can set up 8 or 12 dilution series at the same time. Use 10 mM MgSO₄ to dilute bacteria, use SMG to dilute phages. Be careful about contaminations. Never ever use the diluent directly out of the bottle. Pour some into a petri dish or into a pipetting container and then pipette it from there into the microplate. Add 90 or 180 µL to every well. This is most easily done with a automated multi-channel pipette or with a stepper.

The dilution series is usually a series of steps with a ten-fold dilution in-between. So you add 10 or 20 µL of your sample to the first well (depending on whether there are 90 or 180 µL diluent in the well). Mix properly. Transfer 10 or 20 µL into the next well. Mix proplerly. And so on.

Here are three little hints:

- Exchange pipette tips regularly (see below). Here is why: assume your sample has a concentration of 10⁹ pfu/mL and a tiny droplet (say 0.1 µL) remains inside the tip after the first transfer. If this droplet is washed out after the 7th transfer (at that point you should have around 10 phages in your well), it will add 10⁵ phages to the well. There goes your dilution series.
- Mix well. Pipetting once up and down and then transfer the sample to the next well is not enough. I have never tested this properly, and before I haven't done it, I am simply paranoid about it and do at least 10 strokes with the pipette before I transfer. This is especially important if the shape of the well makes mixing more difficult, e.g. if you use a PCR-plate for your dilution series. In addition, keep in mind that your pipette moves only 10% of the volume, which doesn't allow an efficient mixing.
- Be careful that you don't skip a step in the series. Happens easily. Especially if someone talks to you while you are working (and you are male). If you suspect that it happened, you can check the height of the liquid in your wells to find the one where you pipetted last. In such a case, it is much easier to gauge the height when you work with 180 instead of 90 µL. Another thing that speaks for 180 µL is the simple fact that larger volumes can be transferred more accurately.

Calculating the original concentration

The whole effort is for nothing if you don't manage to calculate how much phages or bacteria you had in your original sample.

- Count the colonies or plaques on your plate or in your spot.
- Calculate the concentration per mL for the dilution step in which you did the counting. E.g. if you count 18 colonies in a spot of 20 µL, the concentration is 900 cfu/mL.
- Go back all the steps to the original sample, and multiply everytime by 10. E.g. if your spot was the 7th, you multiply by 10⁷. So your original sample has a concentration of 9⋅10⁹ cfu/mL.

There are two pitfalls. The first one lurks if you vary the volume that you use for spotting. One day its 5, another day its 10 µL. You can be sure that you don't remember it the next day when you look at your plates. Therefore write it on the plate. The second pitfall lurks if you sometimes spot the original sample as well. If you forget about that, your calculation is off by a factor of 10. To avoid this, write the dilution of the first and last spot on the plate. E.g. if your dilution series goes from 10^-1 to 10^-8, write -1 → -8, if it goes from 10⁰ (undiluted) to 10^-7, write 0 → -7. Or write it on the microplate and leave it on the bench.

You know that something went wrong if…

…your spots of the dilution series never go to zero plaques. What you should see is a single large zone of lysis in the spots with the highest concentration. The further you go down your dilution series, the more discernible single plaques become, and finally the phages are diluted out. If this doesn't happen, you may have a contamination in your diluent. Because the diluent has the same concentration in all spots, you will never reach zero plaques.

…your spots of the dilution series do not show what you actually did. If there are 200 plaques, then 20, then 2, then 0, that's fine. Textbook-example-fine. If its 215, 19, 4, 0, this is also fine. There is always some error. If its 205, 50, 10, 8, 0, something went wrong. Did you set up 90 µL of diluent in all wells but you transferred 20 µL? Did you check at all whether your pipette was set to the right volume? Did you set the volume properly but then you took another pipette? (It all happened already!)

Exchanging pipette tips

It is generally recommend to exchange pipette tips with every dilution step that is performed. This can result in the use of a lot of tips and I was wondering, whether the same tip could be used repeatedly or whether this really messes up the dilution series.

An E. coli stock (JHC510 in 10 mM MgSO₄, OD 2.0, approximately 10⁹ cfu/ml) was diluted over 6 10-fold dilution steps in a 96-well plate. The stock was pipetted into the first well, the following wells contained 180 µL 10 mM MgSO₄. 20 µL were transferred between wells, after every transfer, contents of the well were mixed with 25 pipetting strokes. 5 different pipette tips exchange regimes were used (8 replicates each): every time, every 2^nd, 3^rd, 4^th time, and never. 20 µL of every dilution step was spotted on a plate). Tip exchange (and thus tips used): Every time: 6 tips, every 2^nd time: 3 tips, every 3^rd time: 2 tips, every 4^th time: 2 tips, never: 1 tip.

Figure 1: Estimated number of bacterial cells in the undiluted stock. Every dilution series consisted of 8 replicates. There is no effect of tip exchange regime on the estimated number of bacterial cells in the stock (F_4,35 = 1.62, P = 0.19). There is no significant difference in variances among the regimes (Levene's test, P = 0.85).

It looks like tips don't need to be exchanged at all. There was neither evidence for carry over nor for an increase in variation. However, I recommend to exchange tips every 2nd time. This reduces consumption of tips by 50%. Although the above data don't support this, I expect that errors can occur when tips are exchanged less frequently.