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What is your favorite method for making chemically competent E.coli cells?


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7 replies to this topic

#1 labtastic

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Posted 11 October 2013 - 10:13 AM

What is your favorite way to make chemically competent cells? I commonly do double transformations (for two plasmid expression systems) so I like to make my cells as competent as possible. Plus other folks in the lab are convinced they need commercially prepared competent cells to do their cloning work, which I think is a waste of lab funds considering how much cloning we do. 

 

Historically, I've been quite pleased with Inoue method, but on occasion some batches come out better than others.

 

Can anyone compare that method with, say, the Rubidium chloride method or the CCMB80 method?  I'm thinking of giving either a try for my next batch, but only if there are folks who can testify that it really is better than the Inoue method.

 

 



#2 jerryshelly1

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Posted 11 October 2013 - 10:59 AM

I have tried multiple methods.  My favorite is the Hanahan protocol. Our lab never had any rubidium chloride, but I did scrounge some from an adjacent lab.  I tried the Hanahan protocol side-by-side with rubidium chloride, potassium chloride, and cobalt hexamine chloride.  They all produced negligible transformation efficiencies.

 

The biggest factor is the efficiency of the cells you start with, the media you grow your cells in, the temperature at which you prepare your cells, and the method you handle them. This will drastically affect the efficiency of your cells.

 

I used to work in a similar environment.  Why spend $200.00 for 1mL of competent cells when you could easily make 10mL of competent cells for a fraction of the cost. Comparing my cells to the manufacturer indicated very similar efficiencies. I always try to save money wherever possible.

 

Edit - Incoherent sentence


Edited by jerryshelly1, 11 October 2013 - 11:01 AM.


#3 phage434

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Posted 11 October 2013 - 11:07 AM

The "Hanahan protocol" is very ambiguous. He had about six or seven. Perhaps you could be more specific.



#4 labtastic

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Posted 11 October 2013 - 11:59 AM

I agree with phage...could you be more specific by what you mean with the Hanahan protocol? The top two protocols that google comes up with are different-- one uses rubidium, Mn and Ca, and the other uses cobalt, Mn and Ca.

 

I agree that in this day and age of funding, any place in the lab where you can save a few dollars here and there can add up to thousand dollars over the course of a year, which can make a big difference. Our lab is no exception, and I would really like to convince my lab mates to stop buying commercially prepared cells, even if it means me spending a Saturday morning making a huge batch for the lab. Currently I just make them for myself and they always do the trick for me, but for some reason (poor molecular biology techniques?) they can't make them work to their "satisfaction" so $$'s just get poured down the drain. 


Edited by labtastic, 11 October 2013 - 12:03 PM.


#5 jerryshelly1

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Posted 14 October 2013 - 06:02 AM

Sure.  I will post the one I use when I hit my bench today.



#6 phage434

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Posted 14 October 2013 - 06:25 AM

It would be helpful if  you could include information about their measured competence.

Thanks.



#7 jerryshelly1

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Posted 14 October 2013 - 07:46 AM

Hey guys,

 

Two pasted documents. The original protocol and the modified one. Hope this helps.

 

First:

 

XL-10 and Sure-2 Competent Cell Preparation:

 

Day 1:

·      Make appropriate buffers and Media.

·      Inoculate 2mL of media with frozen cell stock (Starter Culture).

·      Incubate O/N at 37°C.

·      TB Buffer was used for both.

·      SOB+ Media was used

 

Day 2:

·      Add fresh buffers to media.

·      Inoculate 2mL culture into 250mL of media.

·      Shake/Incubate at 25°C until OD400-500 (Smaller OD provides better efficiency).

·      In cold room, Transfer cells to cold 50mL falcon tubes and incubate on ice for 30min.

·      Centrifuge cells at 4000xg for 10min at 4°C.

·      In cold room, decant the supernatant and keep the tubes inverted for several minutes to drain off excess media.

·      Gently resuspend cells in 20mL (per 250mL of appropriate buffer).

·      Incubate on ice for 20min.

·      Centrifuge cells at 4000xg for 10min at 4°C.

·      In cold room, decant the supernatant and keep the tubes inverted for several minutes to drain off excess media.

·      Gently resuspend cells in 10mL (per 250mL of appropriate buffer).

·      Incubate on ice for 20min.

·      Add 350mL of DMSO (per 250mL of cells), dropwise and mix gently.

·      Incubate on ice for 10min.

·      Aliquot appropriate amount of cells to chilled eppendorf tubes.

·      Shock-freeze cells in ethanol dry ice bath

·      Store cells at -80°C.

 

Growth Kinetics:

2hr

·      Sure2 – 0.030

·      XL-10 – 0.005

3hr

·      Sure2 – 0.097

·      XL-10 – 0.034

4hr

·      Sure2 – 0.221

·      XL-10 – 0.051

5hr

·      Sure2 – 0.341

·      XL-10 – 0.090

6hr

·      Sure2 – 0.458 (Harvested)

·      XL-10 – 0.084

 

*At this point it was obvious the XL-10 would take 16+ hrs to grow.  I switched the cells to 18°C and let them grow O/N (7:20pm – 9:00am).

 

22hr

·      XL-10 – 0.536 (Harvested)

 

 

*I checked the efficiency with BME, 1mM CaCl2 and plain at 12 hr.

·      XL-10: 8.1e7

·      XL-10 BME: 2.0e7

·      XL-10 1mM CaCl2: 9.5e8

·      Sure2: 4.6e8

·      Sure2 BME: 1.1e8

·      Sure2 1mM CaCl2: 9.4e7

 

Second:

 

*For Sure-2 cells use NZY+ media.

Protocol for the preparation of competent cells

Andreas Leibbrandt

Source: modified Hanahan procedure after Methods Enzymol. 1991; 204:63-113

 

Buffers to render cells competent:

  • DH5a ® FSB + 5% sucrose
  • XL10-Gold ® FSB or TB (Will use TB buffer, unless I can find HexCo(III)Cl)
  • TOP10 ® CCMB80

            FSB buffer ± sucrose:            10 mM KOAc pH 7.5

                                                100 mM KCl

                                                45 mM MnCl2

                                                10 mM CaCl2

                                                3 mM HexCo(III)Cl  - Cobalt (III) Hexamine Chloride

                                                10% glycerol

                                                [5% sucrose]

                                                pH adjusted to 6.4

 

            TB buffer:                        10 mM PIPES pH 6.7

                                                55 mM MnCl2

                                                15 mM CaCl2

                                                250 mM KCl (Good Substitue)

 

            CCMB80 buffer:            80 mM CaCl2

                                                20 mM MnCl2

                                                10 mM MgCl2

                                                10 mM KOAc pH 7.0

                                                10% glycerol

                                                pH adjusted to 6.4

Material check list:

500 ml of SOB or SOB+ medium (order from IMP media kitchen; add 6.25 ml of 1M MgCl2 and 6.25 ml of 1M MgSO4 prior to use, i.e. SOB+), use ~250 ml per E. coli strain

o   SOB: for CCMB80-competent cells, i.e. Mach1 and TOP10 cells

o   SOB+: for FSB-competent cells, i.e. DH5a and XL10-Gold cells

o   SOB+, 40 mg/ml Cam, 80 mg/ml Tet: for XL10-Gold cells

2 l Erlenmeyer flask with red lid (ask the IMP media kitchen to rinse and autoclave as for cell culture glassware)

0.5 l Erlenmeyer flask (ask the IMP media kitchen to rinse and autoclave as for cell culture glassware)

10 ml Falcon 2059 tube

pre-cooled CCMB80, TB, or FSB buffers

pre-cooled 1.5 ml Sarstedt screw cap tubes (from IMP store)

pre-cooled Eppendorf Combitip 5 ml

pre-cooled Falcon tubes, 50 ml

pre-cooled serological pipettes (5 and 10 ml)

ART200 tips

Vortexer in the cold room

liquid N2 or EtOH/dry ice bath in the cold room

ice basket(s) to incubate cells and transfer them from the centrifuge to cold-room

 

 

DAY 1-2

·      pick 5 single colonies and resuspend by gentle vortexing in 1.5 ml of SOB(+) in a Falcon 2059 tube

o   e.g. from a 10-6 dilution prepared from a frozen stock of competent cells, plated on SOB(+)  agar plates and grown o/n @ 37°C

o   alternatively, scrape off some cells from a frozen glycerol stock and resuspend by gentle vortexing in 1.5 ml of SOB(+)  in a Flacon 2059 tube

·      inoculate in 20 ml of SOB(+)  in a 0.5 l Erlenmeyer flask and grow @ 18°C until the culture becomes turbid

DAY 3-4

·      on the next day, dilute 1:100 in fresh SOB(+)  medium and grow cells to an OD600 of ~0.35-0.6 @ 18°C

o   growth @18°C is very slow, so it might be best to start of ~noon the day before to finish the preparation on the next day

·      transfer cells to 5 cooled 50 ml Falcon tubes, and incubate on ice for ~15-30'

o   optionally: prepare glycerol stock, i.e. cells 1:1 with 60%SOB, 40% glycerol

·      spin down cells @ 4000 rpm for 15' at 4°C

o   don't forget to pre-cool the centrifuge and centrifuge containers

·      in the cold room, decant the supernatant and keep the tubes inverted for several minutes to drain off excess media

·      pool the cells (i.e. 5 Falcon tubes) by resuspending the cell pellets carefully (by gentle vortexing or pipetting) in 1/80-1/85 of the original volume in the respective buffer (i.e. for 250 ml of cells, use 3 ml of buffer)

·      incubate bacteria on ice for 20'

·      for FSB preparations, add 3.5% DMSO (105 µl DMSO/3 ml buffer/ 250 ml SOB(+)) from a freshly thawed aliquot of DMSO to cells, mix by gently swirling the tube and incubate on ice for 10'

o   DMSO is stored @ -20°C; remove an aliquot at the beginning of the procedure since it takes a while to defrost

o   apply DMSO drop by drop to the center of the solution and gently swirl the mixture

·      add the same volume of DMSO as before, mix, and further incubate on ice for 5'

·      aliquot 0.2 ml of DMSO-treated competent cells into pre-cooled 1.5 ml Eppendorf tubes and shock-freeze competent cells in liquid N2 or an EtOH-dry ice bath, store cells @ -80°C

o   aliquot cells by using the Eppendorf Multipette with a pre-cooled 5 ml Combitip

 

 

NZY+ Broth (per Liter):

 

10 g of NZ amine (casein hydrolysate)

5 g of yeast extract

5 g of NaCl

Add deionized H2O to a final volume of 1 liter.  Adjust to pH 7.5 using NaOH.

Autoclave.

Add the following filer-sterilized supplements prior to use:

12.5 ml of 1 M MgCl2

12.5 ml of 1 M MgSO4

20 ml of 20% (w/v) glucose (or 10 ml of 2 M glucose) 

 

 

 

*RbCl and Hexamine Chloride can be substituted with KCl. The efficiencies do not differ substantially. 



#8 phage434

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Posted 14 October 2013 - 12:44 PM

Thank you for the detailed protocol. This is quite similar to the one I developed here, also from Hanahan and some patents, based on CCMB80:

http://openwetware.o...competent_cells






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