This protocol relates to the assay that was made regarding whether 3-methylcatechol has been ingested in our fragments or not:
1. 96 well plate
2. 0.0124g of 3-methylcatechol
3. 5mL of dH2O
4. todE fragment
5. todE + RBS fragment
6. Lysis buffer (negative control)
7. Lysate (positive control)
9. 96 well plate reader
1. Weigh out 0.0124g of 3-methylcatechol. Dissolve it in 5mL of dH2O in a small glass flask. Invert the solution until 3-methylcatechol has fully dissolved in water. (Colour change will occur gradually).
2. In the 96 well plates, pipette 90 uL of todE fragment from 0h, 2h, 4h, 6h, 8h and 24h lysate in duplicates.
3. Similarly, pipette 90 uL of todE + RBS fragment from 0h, 2h, 4h, 6h, 8h and 24h lysate in duplicates.
4. Do not add 10 uL of 3-methylcatechol to step 2 and 3 until all the other wells have been filled.
5. In the next well, pipette 90 uL of the negative control, lysis buffer, into the wells in duplicates. Then, add 10 uL of dH2O.
6. In the next well, pipette 90 uL of the positive control, lysate, of each todE and todE + RBS fragments in duplicates.
7. Pipette 10 uL of 3-methylcatechol to the wells that require it. Immediately after that is done, insert the plate into the 96 well plate reader.
8. Read the plate every minute.
9. Using Excel, graph the growth curve.
Table 1. Illustrates the positions of each solution in each well.
1. 35mg/L agar powder
2. Distilled water
3. Any antibiotics required
4. Ampicillin /Carbenicillin (Red)
6. Kanamycin (Blue)
7. T etracycline (Black)
1. Weigh out the agar powder, 35mg per litre. If you are preparing 500ml, you need 17.5mg etc.
2. Add it to the bottle, add the required amount of distilled water. Mix it by shaking it
3. Autoclave it.
4. It’s best to pick it up from the autoclave while it’s still hot. Have your agar, antibiotics and plates ready.
5. Clean your working bench with ethanol. Use Bunsen burner if you are not working underneath a hood.
6. Wait till the agar is cool enough to touch. If you can’t hold the agar in the bottle comfortably for like 30seconds, then it’s not ready to be poured into plates.
7. Add the antibiotics [if needed]. 1 microlitre per 1ml. For 500ml agar, you need to add 500microlitre.
8. Pour 20ml agar per plate. (500ml makes roughly 25 plates). Don’t put the lid on yet. Just put it half-way on.
9. Let the agar is set, close the lids. Use parafilm around the plates and store then in the 4 degrees fridge upside down.
In order to ensure that our bacteria are competent and able to take up new plasmids, we will need to perform a competency assay.
1. Frozen DNA stock
2. 10 pg/µL pcDNA3.1 aliquoted in 15 µL stocks, frozen at -80°C
3. LB media
4. Bacteria (E.Coli)
Transforming the competent cells:
A) 20 pg DNA (2µL)
B) 100 pg DNA (10 µL)
Recovery stage of transformations
1. Add 900µL LB media to both and incubate 60 min at 37°C
2. Plate 100 µL of both on LB + Ampicillin plates (2 plates for each)
3. Place plates at 37°C overnight
4. Next morning count colonies (colony forming units, cfu)
Determining efficiency of transformation:
1. Good competent cells should be 1x10↑7 to 1x10↑8 cfu/µg
2. Usable competent cells will be 1x10↑6 cfu/µg
T3. he efficiency will be adversely affected by ligations, plasmid size, relaxed (non-supercoiled) state of plasmid DNA.
Figure 1: competent cell equation
BioBrick assembly protocol
Restriction Digest Protocol (link)
Before You Start
Estimated time: 30 min. active, 50 min. incubation
You should keep all materials on ice.
At iGEM HQ we use restriction enzymes from New England Biolabs
1. (1) 8-tube strip, or (3) 0.6ml thin-walled tubes
2. BioBrick Part in BioBrick plasmid (Purified DNA, > 16ng/ul)
4. NEB Buffer 2
6. Restriction Enzymes: EcoRI, SpeI, XbaI, PstI
1. Ice and bucket/container
2. Thermal cycler or heating block
1. Add 250 ng of DNA to be digested, and adjust with dH20 for a total volume of 16ul.
2. Add 2.5ul of NEBuffer 2.
3. Add 0.5ul of BSA.
4. Add 0.5ul of EcoRI.
5. Add 0.5ul of PstI.
There should be a total volume of 20ul. Mix well and spin down briefly.
Incubate the restriction digest at 37C for 30min, and then 80C for 20min to heat kill the enzymes. We incubate in a thermal cycler with a heated lid
Run a portion of the digest on a gel (8ul, 100ng), to check that both plasmid backbone and part length are accurate.
Ligation protocol (Link)
After following our restriction digest protocol (which uses 250ng of DNA) you may follow these steps for ligation.
1. Add 2ul of digested plasmid backbone (25 ng)
2. Add equimolar amount of EcoRI-HF SpeI digested fragment (< 3 ul)
3. Add equimolar amount of XbaI PstI digested fragment (< 3 ul)
4. Add 1 ul T4 DNA ligase buffer. Note: Do not use quick ligase
5. Add 0.5 ul T4 DNA ligase
6. Add water to 10 ul
7. Ligate 16C/30 min, heat kill 80C/20 min
8. Transform with 1-2 ul of product
Illustra plasmidPrep Mini Spin Kit Protocol
Protocol for 1.5 ml and 3 ml culture volumes
1. Harvesting of Bacterial Culture
a. Transfer 1.5 ml from a fresh overnight culture to a microcentrifuge tube. To pellet bacteria, centrifuge at full speed (16 000 × g) in a microcentrifuge for 30 seconds. Discard supernatant and re-centrifuge. Remove any residual supernatant using a pipette. 30 seconds 16 000 x g 15
b. If processing 3 ml culture volumes, repeat step a. Pelleted DNA can be stored at -20°C if necessary
a. Cell re-suspension- Add 175 µl Lysis buffer type 7 to the bacterial pellet and thoroughly re-suspend the pellet.
b. Cell Lysis - Add 175 µl Lysis buffer type 8 and mix immediately by gentle inversion (approximately 5 times) until solution becomes clear and viscous. Allow reaction to happen for 4-5 minutes.
c. Neutralisation - Add 350 µl Lysis buffer type 9 and mix immediately by gentle inversion until the precipitate is evenly dispersed. Gently invert until solution clears
d. Flocculent spin - Centrifuge at full speed (approximately 16 000 × g) for 4 minutes.
e. During centrifugation, for each purification that is to be performed, place one illustra plasmid mini-column in one Collection tube.
3. Plasmid Binding
a. Column lysate loading-Carefully transfer the cleared supernatant to the mini-column (approximately 700 µl). Close the lid of the column gently. Centrifuge at full speed (approximately 16 000 × g) for 30 seconds. Discard the flow-through by emptying the Collection tube.
4. Wash (optional-strain dependent)
a. Wash the column with 400 µl Lysis buffer type 9 and centrifuge at full speed (approximately 16 000 × g) for 30 seconds. Discard the flowthrough.
5. Wash & Dry
a. Add 400 µl Wash buffer type 1 to the column and centrifuge at full speed (approximately 16 000 × g) for 1 minute. Carefully discard flowthrough and the Collection tube.
a. Transfer the illustra plasmid mini column into a fresh microcentrifuge tube and add 50 µl Elution buffer type 4 directly onto the center of the column. Incubate the column for 30 seconds at room temperature. Microcentrifuge at full speed (approximately 16 000 × g) for 30 seconds to recover Link: https://cdn.gelifesciences.com/dmm3bwsv3/AssetStream.aspx?mediaformatid=10061&destinationid=10016&assetid=14878.
Competent cells protocol
This protocol is a variant of the Hanahan protocol Hanahan91 using the CCMB80 buffer for DH10B, TOP10 and MachI strains. It builds on Example 2 of the Bloom05 patent as well. This protocol has been tested on NEB10, TOP10, MachI and BL21(DE3) cells. See OWW Bacterial Transformation page for a more general discussion of other techniques. The Jesse '464 patent describes using this buffer for DH5α cells. The Bloom04 patent describes the use of essentially the same protocol for the Invitrogen Mach 1 cells.
Seed Stock Protocol
Before You Start
You can prepare stocks of your bacteria of interest and store at -80°C to seed future batches of competent cells.
Detergent is a major inhibitor of competent cell growth and transformation. Glass and plastic must be detergent free for these protocols. The easiest way to do this is to avoid washing glassware, and simply rinse it out. Autoclaving glassware filled 3/4 with DI water is an effective way to remove most detergent residue. Media and buffers should be prepared in detergent free glassware and cultures grown up in detergent free glassware.
1. Petri plates with SOB agar
2. Sterile loop
1. Detergent-free, sterile glassware and plasticware (see above)
2. -80°C freezer
3. Incubator or platform with shaker
1. Streak TOP10 cells on an SOB plate and grow for single colonies at 23°C (~16hrs) room temperature works well
2. Pick single colonies into 2 ml of SOB medium and shake overnight (14-16hrs) at 23°C room temperature works well
3. Add glycerol to 15%
4. Aliquot 1 ml samples to Nunc cryotubes
5. Place tubes into a zip lock bag, immerse bag into a dry ice/ethanol bath for 5 minutes This step may not be necessary
6. Place in -80°C freezer indefinitely.
Competent Cell Production Protocol
Before You Start
Detergent is a major inhibitor of competent cell growth and transformation. Glass and plastic must be detergent free for these protocols. The easiest way to do this is to avoid washing glassware with detergent, and simply rinse it out. Autoclaving glassware filled 3/4 with DI water is an effective way to remove most detergent residue. Media and buffers should be prepared in detergent free glassware and cultures grown up in detergent free glassware.
Prechill 250mL centrifuge tubes and screw cap tubes before use.
1. 1. SOB
2. CCMB80 buffer
3. 10 mM KOAc pH 7.0 (10 ml of a 1M stock/L)
4. 80 mM CaCl2.2H2O (11.8 g/L)
5. 20 mM MnCl2.4H2O (4.0 g/L)
6. 10 mM MgCl2.6H2O (2.0 g/L)
7. 10% glycerol (100 ml/L)
8. adjust pH DOWN to 6.4 with 0.1N HCl if necessary
9. adjusting pH up will precipitate manganese dioxide from Mn containing solutions.
10. sterile filter and store at 4°C
11. the slight dark precipitate appears not to affect its function
1. Detergent-free, sterile glassware and plasticware (see above)
2. Table-top OD600nm spectrophotometer
1. Ethanol treat all working areas for sterility.
2. Inoculate 250 ml of SOB medium with 1 ml vial of seed stock and grow at 20°C to an OD600nm of 0.3. Use the "cell culture" function on the Nanodrop to determine OD value.
a. OD value = 600nm Abs reading x 10
b. This takes approximately 16 hours.
c. Controlling the temperature makes this a more reproducible process, but is not essential.
d. Room temperature will work. You can adjust this temperature somewhat to fit your schedule
3. Aim for lower, not higher OD if you can't hit this mark
4. Fill an ice bucket halfway with ice. Use the ice to pre-chill as many flat bottom centrifuge bottles as needed.
a. Transfer the culture to the flat bottom centrifuge tubes. Weigh and balance the tubes using a scale
5. Try to get the weights as close as possible, within 1 gram.
a. Centrifuge at 3000g at 4°C for 10 minutes in a flat bottom centrifuge bottle.
6. Flat bottom centrifuge tubes make the fragile cells much easier to resuspend
7. Decant supernatant into waste receptacle, bleach before pouring down the drain.
a. Gently resuspend in 80 ml of ice cold CCMB80 buffer
b. Pro tip: add 40ml first to resuspend the cells. When cells are in suspension, add another 40ml CCMB80 buffer for a total of 80ml
c. Pipet buffer against the wall of the centrifuge bottle to resuspend cells. Do not pipet directly into cell pellet!
8. After pipetting, there will still be some residual cells stuck to the bottom. Swirl the bottles gently to resuspend these remaining cells
9. Incubate on ice for 20 minutes
10. Centrifuge again at 3000G at 4°C. Decant supernatant into the waste receptacle, and bleach before pouring down the drain.
a. Resuspend cell pellet in 10 ml of ice cold CCMB80 buffer.
11. If using multiple flat bottom centrifuge bottles, combine the cells post-resuspension
a. Use Nanodrop to measure OD of a mixture of 200 μl SOC and 50 μl of the resuspended cells
12. Use a mixture of 200 μl SOC and 50 μl CCMB80 buffer as the blank
13. Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test.
a. Incubate on ice for 20 minutes. Prepare for aliquoting
b. Make labels for aliquots. Use these to label storage microcentrifuge tubes/microtiter plates
14. Prepare dry ice in a separate ice bucket. Pre-chill tubes/plates on dry ice.
15. Aliquot into chilled 2ml microcentrifuge tubes or 50 μl into chilled microtiter plates
a. Store at -80°C indefinitely.
16. Flash freezing does not appear to be necessary
a. Perform test transformations to calculate your competent cell efficiency
b. Thawing and refreezing partially used cell aliquots dramatically reduces transformation efficiency by about 3x the first time, and about 6x total after several freeze/thaw cycles.
c. Good cells should yield around 100 - 400 colonies
d. Transformation efficiency is (dilution factor=15) x colony count x 105/µgDNA
We expect that the transformation efficiency should be between 1.5x108 and 6x108 cfu/µgDNA
5x Ligation Adjustment Buffer
Intended to be mixed with ligation reactions to adjust buffer composition to be near the CCMB80 buffer
1. KOAc 40 mM (40 ml/liter of 1 M KOAc solution, pH 7.0)
2. CaCl2 400 mM (200 ml/l of a 2 M solution)
3. MnCl2 100 mM (100 ml/l of a 1 M solution)
4. Glycerol 46.8% (468 ml/liter)
5. pH adjustment with 2.3% of a 10% acetic acid solution (12.8ml/liter)
6. Previous protocol indicated the amount of acetic acid added should be 23 ml/ litre but that amount was found to be 2X too much per tests on 1.23.07 --Meagan15:50, 25 January 2007 (EST)
a. water to 1 litre
7. autoclave or sterile filter
8. Test pH adjustment by mixing 4 parts ligation buffer + 1 part 5x ligation adjustment buffer and checking pH to be 6.3 - 6.5
9. Use of the ligation adjustment buffer is optional.