Yes, volumes matter. Remember, your competent cells are spheroplasts that are not protected by their cell wall. Therefore, they are very sensitive to osmotic stresses. If you had used an isotonic solution, maybe this phenomenon would be less apparent.

Another factor here is that at very low concentrations, loss of DNA due to adsorption to pipet tips or microfuge tubes can become a problem. Since it is difficult to quantify such low amounts of DNA, we can't really be sure you had equivalent total plasmid in your inputs.

Yuji and Juan also raise a legitimate concern about residual DEPC. But I wouldn't be surprised if your results reproduced with non-DEPC water.

The takeaway here is that if you are running QC on competent cells, try to keep the volume to

SImple question, the DEPC treated water has also been autoclaved to inactivate the DEPC? maybe the DEPC itself is interfeering with the trasnformation or even having a toxic effect on the cells (since the more water you used the less colonies you had).

Hi Elia,

Yes, the DEPC-treated water has been autoclaved as well.

Did you mix the10 uL volume by pipetting? It's been a while but I seem to remember the competent cells being susceptible by shearing and agitation.

Just needed to be sure. What is the volume of cells in which you electroporate? Usually we prepare aliquotes of 45ul, if you do the same then adding 10ul of water this migth give an osmotic shock to the cells. But it's just my theory, I never had such experience. This migth be a possible explanation the reason why we prepared our protocol to use at best 4ul (the macimum volume I ever used on 45ul of cells).

My understanding has always been that the volume that the DNA is in has to be low. My guess is that the excess water is effecting transformation. The addition of otherwise de-ionized water may cause osmotic stress or dilute salts that make the bacteria chemically competent. It also lowers the concentration of the DNA in the transformation. However, as little as 0.1ng of DNA is needed for transformation of plasmids that are ~5kbp and I have used much less if it is pure supercoiled DNA. I also think that with XL1 blue it is important that heat shock is performed for only 45sec. I have noticed that increasing heatshock time with XL1 blue will decrease transformation efficacy.

@Aaron: You're right, mixing by pipetting isn't a good idea and I didn't do it. I only tapped the samples the way I always do it. The treatment was identical for both samples.

@Elia: I use 200 µl of chemically competent cells. I know that you're not supposed to add more than 10% of the total volume. Nevertheless, I didn't expect such a profound difference between adding 1 µl or 10 µl (both should be fine in 200 µl).

@Ted: I can only assume that the lowered concentration is is responsible for that effect. Nevertheless, the effect is very profound and I am just surprised to see that.

I used XL2 blue cells and with 1 min heat shock I achieved a transformation efficacy of 1.5 x 10(7)/µg pUC19. It may be that 45 s works better, but 1 min works quite well. I can try to see if I need to optimize the heat shock time.

Hi, Chris.

Similar problem occured when I transfered 2 and 10 μl to self-made DH5α. More colonies observed for 2μl tube. But it happened only once, I didn't repeat the experiment to confirm it. Have you repeated the experiment and got the same result? At that time, I thought it was caused by a too hot glass spreading rod, which might kill the bacteria when spreading the plate.

From the attatched file, it seems that transformants are possitive related to concentration of plasmid DNA.

http://www.itescam.edu.mx/principal/sylabus/fpdb/recursos/r72068.PDF

Why don't you use just pure water autoclaved?

I guess residual of DPEC can inhibit transformation efficiency.

Yes, I agree with this last answer, What is inhibiting transformation is the residual of DPEC. Just dilute your plasmid DNA in MilliQ water autoclaved.

Transformation efficiency besided many factors also depends upon how concentrated the DNA is, and how diluted the other buffers/ water is - in which DNA was present. up to my experience the DNA that is transformed should be more then 95% pure and as concentrated as possible. Your results are example for it.

Yes, volumes matter. Remember, your competent cells are spheroplasts that are not protected by their cell wall. Therefore, they are very sensitive to osmotic stresses. If you had used an isotonic solution, maybe this phenomenon would be less apparent.

Another factor here is that at very low concentrations, loss of DNA due to adsorption to pipet tips or microfuge tubes can become a problem. Since it is difficult to quantify such low amounts of DNA, we can't really be sure you had equivalent total plasmid in your inputs.

Yuji and Juan also raise a legitimate concern about residual DEPC. But I wouldn't be surprised if your results reproduced with non-DEPC water.

The takeaway here is that if you are running QC on competent cells, try to keep the volume to

Many thanks to everyone for the helpful discussion. I appreciate your help.

I agree with Daniel. I wouldn't use more than 5 ul per 50 ul of competent cells. I have this experience: a number of transformants doesn't correlate linearly with DNA concentration when different volumes of DNA are used.

I fully agree with the collegues Juan Ayala and Yuji Morita: the residual of DEPC decreases the transformation rate. Moreover , from my experience, the efficacy sometimes dependent from the plasmid concentration

The other issue is that if you purified the DNA using a spin column method, the last wash step typically contains alcohol. If you are not careful, some of the wash buffer can be left at the bottom of the the spin column where the seal is (hard to see but you can use a pipette to pull up 5-10 uL of wash solution). Then when you add water to elute your DNA it gets contaminated with the alcohol in the wash buffer. That contamination of your DNA will affect the cells, and the more of the DNA solution you add to the cells, the more alcohol and other contaminants the cells will see and that will likely reduce the transformation efficiency.

On the other hand, if you used too many cells during the lysis step of the mini prep, the final DNA will not be very clean and that will decrease the transformation efficiency as you increase the volume of DNA you add to the cells.

You want to keep the volume of DNA less than 10% of the volume of the cells. Preferentially as little as possible.