Aug 11, 2022
Transforming E. coli (Instructor protocol)
This protocol is a draft, published without a DOI.
- 1University of Wisconsin - Stout
Protocol Citation: Brian Teague 2022. Transforming E. coli (Instructor protocol). protocols.io https://protocols.io/view/transforming-e-coli-instructor-protocol-ce44tgyw
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: August 10, 2022
Last Modified: August 11, 2022
Protocol Integer ID: 68476
Keywords: competent, E. coli, outgrowth, selection
Abstract
This is the instructor protocol for
Protocol
NAME
Transforming E. coliCREATED BY
Brian Teague
Setup for this lab can be pretty intensive if you're starting from scratch. It requires:
- Competent E. coli
- LB-agar + kanamycin plates
- SOC outgrowth media
Notably, commercial competent E. coli are really expensive. We use the Zymo kit, which brings the cost down to ~50 cents per transformation. This protocol makes about 100 transformations, and the cells are competent enough for subcloning, regular restriction-and-ligation cloning, and Golden Gates.
In an attempt to make this work more widely accessible, I have been extremely verbose in these protocols. If you are familiar with recombinant DNA work, you likely have your own protocols for making chemically competent E.coli, pouring plates, and making SOC -- use those!
Materials
Equipment
- Autoclave
- several 1L and 250 ml bottles
- A 250 ml baffled flask
- Shaking incubator
- Spectrophotometer & micro cuvettes
- Nanodrop (or similar instrument for measuring DNA concentration, such as a Qbit or Dynaquant)
- Refrigerated swinging-bucket or high-speed centrifuge
- Cold room (not strictly required but highly recommended)
- -80 °C freezer
Materials and Reagents
- Microcentrifuge tubes, sterile
- 15 ml and 50 ml conical centrifuge tubes
- Petri dish, 10cm, polystyreneFisher ScientificCatalog #FB0875712
- KanamycinResearch Products International (rpi)Catalog #K22000-25.0 (or kanamycin from another vendor, or kanamycin solution at 50 mg/mL )
- 10 mL syringesBD BiosciencesCatalog #BD 309695
- 0.2 µm syringe filterCorningCatalog #CLS431212
- TryptoneFisher ScientificCatalog #BP1421-500
- BD Bacto™ Yeast Extract BD BiosciencesCatalog #212750
- Sodium ChlorideFisher ScientificCatalog #S271
- Potassium chlorideSigma AldrichCatalog #P9333
- Magnesium chloride hexahydrateSigma Aldrich
- Magnesium sulfate heptahydrateSigma AldrichCatalog #M2773
- α-D-GlucoseSigma AldrichCatalog #158968 solution, 40 Mass Percent (autoclaved or filter-sterilized)
- Agar, bacteriological gradeAmrescoCatalog # J637
- Magnesium sulfate heptahydrateSigma AldrichCatalog #M2773
- LB Broth
- Mix & Go! E.coli Transformation Buffer SetZymo ResearchCatalog #T3002
- Monarch Plasmid Miniprep KitNEBCatalog #T1010 (or equivalent miniprep kit)
- Monarch DNA Elution Buffer - 25 mlNew England BiolabsCatalog #T1016L
- Glass beads 5 mmVWR ScientificCatalog #26396-596
Protocol materials
BD Bacto™ Yeast Extract Becton Dickinson (BD)Catalog #212750
In Materials and 2 steps
Magnesium sulfate heptahydrateMerck MilliporeSigma (Sigma-Aldrich)Catalog #M2773
In Materials, Materials and 2 steps
10 mL syringesBecton Dickinson (BD)Catalog #BD 309695
Materials, Step 4
Monarch® Plasmid Miniprep Kit New England BiolabsCatalog #T1010
Step 42
Glass beads 5 mmVWR InternationalCatalog #26396-596
Materials, Step 45
Sodium ChlorideFisher ScientificCatalog #S271
In Materials and 2 steps
0.2 µm syringe filterCorningCatalog #CLS431212
Materials, Step 4
TryptoneFisher ScientificCatalog #BP1421-500
In Materials and 2 steps
Monarch DNA Elution Buffer - 25 mlNew England BiolabsCatalog #T1016L
Materials, Step 44
Magnesium chloride hexahydrateMerck MilliporeSigma (Sigma-Aldrich)
Materials, Step 8
Potassium chlorideMerck MilliporeSigma (Sigma-Aldrich)Catalog #P9333
Materials, Step 8
KanamycinResearch Products International Corp (RPI)Catalog #K22000-25.0
Materials, Step 2
α-D-GlucoseMerck MilliporeSigma (Sigma-Aldrich)Catalog #158968
Materials, Step 11
LB Broth
Materials, Step 24
Monarch Plasmid Miniprep KitNEBCatalog #T1010
Materials
Petri dish, 10cm, polystyreneFisher ScientificCatalog #FB0875712
In Materials and 2 steps
Mix & Go! E.coli Transformation Buffer SetZymo ResearchCatalog #T3002
In Materials and 2 steps
Safety warnings
Several of these chemicals are moderately hazardous, particularly the ones in the miniprep kit. Wear appropriate PPE, including gloves, safety glasses and a lab coat.
This protocol involves the creation or manipulation of genetically modified organisms. Make sure cultures and contaminated plastics are disposed of only after inactivating the GMOs, such as by autoclaving or treating with bleach.
Prepare kanamycin stock solution
Prepare kanamycin stock solution
Prepare 9 empty microcentrifuge tubes in a rack.
Weigh 0.5 g of KanamycinVWR InternationalCatalog #K22000-25.0 into a 15 ml conical centrifuge tube.
Add 10 mL deionized water to make a stock solution with a concentration of 50 mg/mL . Vortex to dissolve.
Mount a 0.2 µm syringe filterVWR InternationalCatalog #CLS431212 on a 10 mL syringesVWR InternationalCatalog #BD 309695 . Pull the plunger out of the back and pour the kanamycin solution in.
Holding the syringe filter over the first microcentrifuge tube, insert the plunger back into the syringe. Squeeze the syringe to filter the kanamycin into the waiting tubes. Put about 1.2 mL into each tube. You don't have to be precise, but make sure there's at least 1 mL in each.
Use immediately or store at -20 °C .
Making SOB and SOC outgrowth media
Making SOB and SOC outgrowth media
In a 250 ml bottle, add approximately 200 mL deionized H2O.
Add to the bottle:
- 5 g TryptoneVWR InternationalCatalog #BP1421-500
- 1.25 g BD Bacto™ Yeast Extract VWR InternationalCatalog #212750
- 0.145 g Sodium ChlorideVWR InternationalCatalog #S271 (or 0.5 mL of a 5 Molarity (M) solution)
- 0.0475 g Potassium chlorideVWR InternationalCatalog #P9333 (or 125 µL of a 1 Molarity (M) solution)
- 2.5 mL 1 Molarity (M) solution of Magnesium chloride hexahydrateVWR International
- 2.5 mL 1 Molarity (M) solution of Magnesium sulfate heptahydrateVWR InternationalCatalog #M2773
Add deionized water to a final volume of 250 ml. (You can eyeball it -- no need to dirty a graduated cylinder).
Autoclave at 121 °C on a liquid cycle for 00:30:00 . (This media is SOB - "Super Optimal Broth")
30m
Using good sterile technique, make 5 mL aliquots into 15 ml conical centrifuge tubes, 1 per 4 people. To each aliquot, add 50 µL 40 Mass / % volume α-D-GlucoseVWR InternationalCatalog #158968 . (These aliquots are SOC - "Super Optimal Broth, Catabolic")
Note
Make several extra! These become contaminated really easily.
Pour LB-agar + kanamycin plates
Pour LB-agar + kanamycin plates
Fill a 1 liter screw-cap bottle with approximately 900 mL of deionized water.
Add:
- 5 g BD Bacto™ Yeast Extract VWR InternationalCatalog #212750
- 10 g TryptoneVWR InternationalCatalog #BP1421-500
- 10 g Sodium ChlorideVWR InternationalCatalog #S271
- 20 g Agar, bacteriological gradeVWR InternationalCatalog # J637
Add water to a total volume of 1 L (eyeballing is OK, no need to dirty a graduated cylinder). Cap and shake to mix.
Note
Make sure you get all of the powder off of the bottom of the bottle. It doesn't have to be completely dissolved, just resuspended.
Loosen the cap and autoclave at 121 °C for 30 minutes on a liquid cycle.
Swirl to mix well, then cool the bottle to at or below 60 °C . You can do this in a water bath, or by swirling under a running cold water tap.
Note
If the media is too hot when you add the antibiotic, it will break down.
Note
My old grad student mentor used to say "if you can hold your hand against it for 60 seconds, it's cool enough." Or you could use an infrared thermometer gun. It's useful for the bottle to be cool enough to hold bare-handed, though!
Optional: pour several plates without any antibiotic. They'll be useful below!
Add 1 mL of 50 mg/mL kanamycin solution and swirl to mix well.
Option A: Pour ~15 ml of molten media into each Petri dish, 10cm, polystyreneVWR InternationalCatalog #FB0875712 , enough to cover the bottom . If you make 1 L of media, you'll use about 2 sleeves (25/sleeve).
Option B: Using a 25 ml pipette and a pipettor, pipette ~15 ml of molten media into each Petri dish, 10cm, polystyreneVWR InternationalCatalog #FB0875712 , enough to cover the bottom . If you make 1 L of media, you'll use about 2 sleeves (25/sleeve).
Leave the plates Overnight on the bench to cool.
Put the petri dishes back in their plastic bags and store inverted at 4 °C . Plates are good for at least 3 months.
Making chemically competent E. coli
Making chemically competent E. coli
Two days before the prep, strike out the E. coli cloning strain (from a freezer stock) on an LB-agar plate (no antibiotics!). Incubate at 37 °C Overnight .
30m
The afternoon before the prep, pick a colony off of the plate and start an overnight culture in 5 mL LB BrothVWR International (in a round-bottomed test tube.) Shake 200 rpm, 37°C Overnight
30m
Transfer 50 ml of SOB (above) to a 250 ml baffled flask.
Add 0.246 g Magnesium sulfate heptahydrateVWR InternationalCatalog #M2773
Note
No, this isn't sterile -- but the culture won't be growing long enough for it to be a problem.
Add 0.5 mL of the overnight culture.
Fold a piece of aluminium foil over the mouth of the flask. Shake 200 rpm at room temperature until the OD600 of the culture reaches 0.4-0.6. This usually takes about 3 hours.
Note
Check the culture every hour until the OD600 reaches 0.2, then ever 30 minutes until it is between 0.4 and 0.6. Do not overgrow the cells; if they are overgrown, throw out the culture and begin again from step 27.
While the culture is growing:
- Chill a swinging-bucket or high-speed centrifuge to 4 °C
- Load 96 microcentrifuge tubes into a 96-position tube rack and place them in the cold room to chill (or on ice). Tent a paper towel over them to keep out contaminating microbes
- From the Mix & Go! E.coli Transformation Buffer SetVWR InternationalCatalog #T3002 kit, mix 2.5 ml of 2X Wash Buffer and 2.5 ml Dilution Buffer to prepare 5 ml of Wash Buffer in a conical centrifuge tube. Keep on ice.
- From the Mix & Go! E.coli Transformation Buffer SetVWR InternationalCatalog #T3002 kit, mix 2.5 ml of 2X Competent Buffer and 2.5 ml Dilution Buffer to prepare 5 ml of Competent Buffer in a conical centrifuge tube. Keep on ice.
- Pre-chill a 50 ml conical centrifuge tube on ice.
When the culture has reached an OD600 of between 0.4 and 0.6, transfer the baffled flask to an ice bucket and mound up the ice around the flask. Chill for 10 minutes.
Note
From this point on, everything must be kept as cold as possible!
Transfer the culture from the 250 ml baffled flask to the 50 ml pre-chilled conical centrifuge tube. Centrifuge in a pre-chilled swinging-bucket centrifuge (or high-speed fixed-angle centrifuge) 2000 x g, 4°C, 00:10:00 .
Note
Move the conical from the ice bucket, to the centrifuge, and back to the ice bucket.
10m
In the cold room (if available), decant the media back into the baffled flask, then invert the 50 ml conical onto a paper towel for a minute to let the media drain away.
Resuspend in 5 ml 1X Mix&Go Wash Buffer by gentle votexing (ie, on a setting of 3-4). Be patient, it will take a few minutes for the cells to resuspend at this speed. Put the resuspended cells back on ice.
Note
If you don't have a cold room, alternate between vortexing and incubating on ice. Remember, cold is key!
Centrifuge in a pre-chilled swinging bucket (or high-speed fixed-angle) centrifuge 2000 x g, 4°C, 00:05:00
5m
In the cold room, decant the wash buffer, then invert the 50 ml conical onto a paper towel for a minute to let the wash buffer drain away.
Resuspend the pellet in 5 ml 1X Mix&Go Competent Buffer by gentle votexing (ie, on a setting of 3-4). Be patient, it will take a few minutes for the cells to resuspend at this speed. Put the resuspended cells back on ice.
Pipette 50 ul aliquots into the prepared microcentrifuge tubes.
Note
An electronic pipettor or a repeat pipettor can be a real time-saver here!
Optional but highly recommended - snap-freeze the cells in liquid nitrogen.
Transfer the tubes to a -80°C freezer, trying to minimize the time between cold room and freezer.
Note
Competent cells prepared this way last for at least a year with no practical decrease in transformation efficiency.
Prepare the transformation control
Prepare the transformation control
Two days before: strike out the E. coli harboring the YTK96 plasmid on a LB agar + kanamycin plate
The day before: Pick one colony of the YTK96-harboring E. coli into 5 ml LB broth + 50 ug/ml kanamycin.
Miniprep the plasmid, using Monarch® Plasmid Miniprep Kit VWR InternationalCatalog #T1010 or comparable.
Analyze the concentration and purity of the miniprep using a Nanodrop.
Prepare transformation controls by diluting the YTK96 miniprep to a concentration of 1 ng/ul in Monarch DNA Elution Buffer - 25 mlVWR InternationalCatalog #T1016L or similar.
Note
I generally make this easy - if the concentration is, say, 67 ng/ul, then I put 67 µL of elution buffer in a tube and add 1 µL of miniprep.
Note
Prepare several tubes of control plasmid.
Miscellaneous preparation
Miscellaneous preparation
Pour the Glass beads 5 mmVWR InternationalCatalog #26396-596 into several 15 ml conical centrifuge tubes.
Note
Depending on the bottle, it may be easier to pour them into a 50 ml tube first, then into 15 ml tubes.
Instructor Tips & Common Student Errors
Instructor Tips & Common Student Errors
Instructor Tips
- Do not decrease the incubation times, especially for the outgrowth. I tried decreasing the outgrowth time to 30 minutes, once, and every single transformation failed. Kanamycin is not like ampicillin -- it doesn't just inhibit growth, it actually kills the cells. So they need to be expressing the resistance gene before they are challenged with the antibiotic.
- I recommend testing the competent cells to make sure they are actually competent before handing them to your students.
- Especially the first time, a two-hour lab may not be enough time for the 90 minutes of (total) incubation. I will often instruct students to label and prepare their plates with beads and leave them on the bench for me -- and then, after their incubations are done, I'll plate their cells for them. Timing isn't important here -- I've seen successful transformations after even 3 hours of outgrowth.
- Make sure the water bath and incubator are turned on well before lab starts -- they take a while to come to temperature!
- If you're using a dry bath instead of a big water bath, fill the dry bath's wells with water to increase the efficiency of heat transfer.
- If you don't have an incubating shaker, it's not the end of the world -- a 37 °C incubator is probably fine.
Common Student Errors
- Didn't check the SOC and used a contaminated tube.
- Added the positive control AND their ligation to the competent E. coli.