Hey there,

There are tons of protocols available. I attach one that is simple (chemically competent, not electroporation) and works well, and should give you 10^6 to 10^7 colonies/ug plasmid, enough for standard cloning approaches. If you need supercompetents you should go for fancier buffers with cobalt, and you could change the strain.

If you are making electrocompetents with some protocol someone else will provide, remember that the key step is the wash: make sure it's thorough, to minimise salt carryover, prevent arcing in the electroporator and maximise efficiency.

The key step in preparation of either chemically or electro-competent cells is to keep everything chilled (buffers, and cells once you start doing the washes), and flash-freeze them in either pre-cooled tubes on dry ice, or pre-cooled tubes on ice that go straight away to liquid nitrogen.

Also, make sure whether your problem is:

• the cells: assess them with a small, known plasmid, as recommended on the protocol I attach.
• the plasmid: some plasmids (especially big ones) will give lower efficiencies.
• the cloning: perhaps your cloning is not efficient

You can check the cloning:

• the plasmid backbone and insert are correct. If you are doing a digestion, you may run them on a gel if you expect changes in band size.
• the plasmid is being created. Use PCR, with primers flanking the cloning points, along a positive control. For instance, if the insert is not too big, you can amplify empty plasmid (positive control) and plasmid after cloning (the band size should increase with the insert).

Just give it a thought, because problems can be anywhere on the protocol.

Good luck,

Ruben

Hi Ruben,

Thank you so much for this detailed answer. I am doing electroporation, and my plasmid is big. Cells are XL1blue and plasmid is pFNOM6-SH2-library (4000 bp). For preparation of electrocompetent cells I am washing the cells with chilled autoclaved dd-H2O and chilled 10% glycerol for 3 times, and I collect them into 10% glycerol then proceeding with electroporation.

Hello Isin.

I've made competent cells of XL1 a lot of times with this protocol, and always worked for me. I've even succesfully electroporated plasmids of 7kb after a clonning with this protocol.

I hope it will work for you!!

Materials:

- Autoclaved milliQ water

- Autoclaved 10 % glycerol.

- Centrifugue bottles (I use 250 ml ones)

- Eppendorfs and tips

- Refrigerated centrifuges (250ml bottles and eppendorf tubes)

EVERYTHING SHOULD BE ICE COLD

1. Pour 100ml culture in 2 centrifuge bottles. Keep them on ice 15 minutes.

2. Centrifuge 5000g, 15 min

3. Resuspend each pellet in 50 ml ice cold milliQ water. Don't use tips. Shake the bottle mannually over the ice.

4. Centrifuge 6000g, 15 min

5. Resuspend each pellet in 25 ml ice cold milliQ water.

6. Centrifuge 7000g, 15 min.

7. Resuspend each pelelt in 500ul 10% glycerol. Combine both pellets in the same eppendorf tube.

8. Centrifuge eppendorf tube 7000g, 5 min.

9. Resuspend in 250ul 10% glycerol. Prepare aliquots of 50ul. Use them fresh or store them at -80ºC

Hi Nuria,

Thank you so much for the procedure. I will try that for sure, I have been re suspending with tips and also I was using 10%glycerol for my wash steps. I have anther question regarding that, to you, do you do 10^-2 10^-4 10^-6 dilutions and plating them onto plates to observe the efficiency?

Thanks a lot,

Hi Isin.

For me, after eletroporation, I add 500ul LB very quickly. After 1 hour of incubation for expression, I plate 100, 150 and 250ul in selective plates.

To be honest, I've never checked the efficiency because it always worked for me!

hi

this the protocol I have used

Preparation of electro-competent cells

An overnight colony was grown on 5ml LB at appropriate temperature and 200 rpm agitation with or without antibiotic according to the culture. Then 1 ml of this culture was inoculated into 100 ml of LB and incubated in the previous conditions until OD600= 0.3-0.5. The culture then was divided into two 50 ml Falcon tubes and chilled on ice for 15 min then centrifuged at 4000 rpm, 4°C for 10 min. The pellets were washed twice with 25 ml of 10% cold glycerol. The pellets were washed again with 12.5 ml of ice-chilled glycerol and centrifuged as before. The washed pellets were suspended on 0.2 ml of 10% ice-chilled glycerol and 50 µl aliquots stored at - 70°C.