GenProtocols

From 2008.igem.org

Contents

QIAprep Spin Miniprep Kit

This protocol is designed for the purification of up to 20 ug high-copy plasmid DNA from 1-5 ml overnight E. coli culture in LB medium.

Procedure

  1. Resuspend pelleted bacterial cells in 250uL Buffer P1 and transfer to a microcentrifuge tube.
  2. Add 250 uL Buffer P2 and mix thoroughly by inverting the tube 4-6 times.
  3. Add 350 uL Buffer N3 and mix immediately and thoroughly by inverting the tube 4-6 times.
  4. Centrifuge for 10 min at 13,000 rpm in a table-top microcentrifuge.
  5. Apply the supernatant to the QIAprep spin column by pipetting
  6. Centrifuge for 30-60 s. Discard flow-through.
  7. Wash QIAprep spin column by adding 0.75 ml BUffer PE and centrifuging for 30-60s.
  8. Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.
  9. To elute DNA, place the QIAprep column in a clean 1.5 ml microcentrifuge tube. Add 50 uL Buffer EB to the center of each QIAprep, let stand for 1 min, and centrifuge for 1 min.


Ligation protocol, using Roche Rapid Ligation Kit

Brief Protocol

Mix:

  • 5x DNA dilution buffer – 2uL
  • Vector (cut and CIP treated) – 0.25-1uL
  • Insert DNA – 0.5-6.5 uL
  • ddH2O – to final volume of 10uL


Vortex and quick centrifuge above mixture, then add:

  • 10uL of 2X rapid ligation buffer
  • 1 uL ligase

Vortex and quick centrifuge again. Hold at room temp ~20 minutes, then use 5uL of the ligation reaction to transform 50uL of chemically competent E. coli (eg, TOP10 or DH5-alpha).

Protocol Notes

Reagents

Make sure buffers are completely thawed before use. The 2x rapid ligation buffer contains a reducing agent that looks like white flakes when thawed. These white flakes must be completely dissolved back into the buffer before use. Keep the ligase enzyme at -20 degrees until just before you are ready to add it, and return it to the freezer promptly. Minimizing time outside of the freezer will preserve the activity of the enzyme.

DNA quantities to use

The ratio of insert DNA to vector is very important. Ideally, you want 10 units of insert to each 1 unit of vector. To estimate this, use the Nanodrop to quantitate the amount of vector and insert you have. Note, the Nanodrop will give you concentrations in nanograms, so you will need to roughly convert this to a ratio based on the size of the insert and the vector.

Example: Vector DNA is 4000 base pairs, and you have 100ng per uL concentration. You use 0.5 uL (50ng) of vector. Your insert is 800 base pairs, and you have 50ng per uL. Your vector is 5 times as large as your insert, which means that on a per-nanogram basis, there are 5 times as many individual units of insert per nanogram in your insert sample than in your vector sample. So, you should use only double the nanogram quantity of insert (2uL) in this reaction (5x more units per ng X 2uL = 100ng = 10x insert compared to vector). As you can see, this is a very rough calculation, but it works pretty well.

Troubleshooting

What to try if your ligation isn’t working (in this order). Foremost among these is to ask your TFs for help and advice. Ask for help sooner rather than later to avoid headaches!

  1. Repeat the ligation. Set up three reactions, one using 0.5uL of insert, one with 2uL of insert and one with the maximal volume the reaction will allow (7-7.5uL insert). Do all of these reactions in parallel. Sometimes, the above mentioned rough quantitation scheme doesn’t work, so it is best to try a range of insert to vector ratios.
  2. Review all of your digestion steps – are you sure you cut the vector and the insert with the correct enzymes? Did you let it digest for long enough? Did you CIP treat the vector after the digest by before the purification? (Many people don’t use CIP on their cut vectors, but I find it increases the success of your cloning endeavours enormously). If you have any doubts about the digestion steps, do them again and then repeat the ligation with three quantities of insert, as described above.
  3. Rarely, enzymes do go bad (perhaps a labmate unintentionally left the enzyme at room temp for too long and then put it back). Try a new kit.
  4. Still not working? Send the parts you are trying to clone for sequencing. Often, DNA obtained from other scientists or from the parts registry is contaminated or just plain wrong. You may not be cloning what you think you are cloning.


Bacterial Transformation Protocol

Chemically competent cells

To make chemically competent cells

Day 1:

  1. Grow 5mL culture of bacteria in LB media overnight, 37° C , shaking

Day 2:

  1. Inoculate fresh 100mL LB culture with 1mL bacteria from 5mL culture
  2. Grow culture at 37°, shaking, to an OD600 of 0.2-0.4 (takes 1-3h.)
    Perform the remainder of the protocol in the cold room OR take care to make sure cells remain cold!!!
  3. Decant culture into two 50mL falcon tubes and chill on ice 15 minutes.
  4. Spin for 15 minutes at 2500 RPM, 4°C
  5. Remove supernatant and wash pellets in 50 mL ice cold 100mM MgCl2 (add 25 mL per tube and combine tubes)
  6. Spin for 15 minutes at 2500 RPM, 4°C
  7. Remove supernatant and resuspend pellet in 50mL ice cold 100mM CaCl2
  8. Incubate on ice 30 minutes
  9. Spin for 15 minutes at 2500 RPM, 4°C
  10. Remove supernatant and gently resuspend pellet in 10 mL ice cold 100mM CaCl2 + 15% glycerol (v/v).
  11. Incubate on ice 30 minutes.
  12. Aliquot 100uL to 0.6mL tubes and store at -80°C. (makes ~100 aliquots)


To transform chemically competent cells

  1. Soak the spots in 5 µL of the warmed TE for 20 minutes. This allows the maximum concentration of DNA in solution. Start thawing the competent cells on wet crushed ice.
  2. Chill labeled 2 ml conical bottom tubes on wet ice. Add 2 µL of DNA in TE and 50 µL of thawed TOP10 competent cells to the tubes. In our experience, these volumes have the best transformation efficiency. The 2 ml tubes allow better liquid movement during incubation. Extra eluted DNA may be held at least several weeks frozen or at refrigerator temperature.
  3. Hold the DNA and competent cells on ice for 30 minutes. This improves transformation efficiency by a significant amount.
  4. Heat shock the cells by immersion in a pre-heated water bath at 42ºC for 60 seconds. A water bath is important to improve heat transfer to the cells.
  5. Incubate the cells on ice for 2 minutes.
  6. Add 200 μl of SOC broth (check that this broth is not turbid, which would indicate previous contamination and bacterial growth). This broth should contain no antibiotics.
  7. Incubate the cells at 37ºC for 2 hours while the tubes are rotating or shaking. We have found that growth for 2 hours helps in transformation efficiency, especially for plasmids with antibiotic resistance other than ampicillin.
  8. Label an LB agar plate containing the appropriate antibiotic(s) with the part number, plasmid, and antibiotic resistance. Plate 250 µl of the incubated cell culture on the plate.
  9. Incubate the plate at 37ºC for 12-14 hours, making sure the agar side of the plate is up. If incubated for too long the antibiotics, especially ampicillin, start to break down and un-transformed cells will begin to grow.


For electrocompetent cells

To make and transform electrocompetent cells

  1. Grow bacteria to log phase (OD 0.4-0.6)
  2. Aliquot 1mL of culture to a 1.5mL epi tube. Make as many aliquots as you need for the number of plasmids you need to transform
  3. Centrifuge aliquots at 12,000g for 1 minute
  4. Remove media and wash with 0.33 volumes (330μL) of 1M sorbitol
  5. Centrifuge at 12,000g for 1 minute
  6. Remove wash and resuspend in 0.05 volumes (40μL) 1M sorbitol.
  7. Place tubes in ice and use within 15 minutes.
  8. Add 100-500ng of plasmid DNA to the tube
  9. Electroporate at 0.55kV
  10. Allow to recover for 1-2 hrs at 30 degrees C

Restriction Digest, using [http://www.neb.com/nebecomm/tech_reference/restriction_enzymes/default.asp New England Biolabs] (NEB) Enzymes

  • Pick a volume where it will be easy to calculate the quantities of 10x and 100x buffers to add and write out a quick recipe (See samples).

(You can add a little water or EB buffer to make this easy, without hurting the reaction)

  • Pipette the DNA to cut into a 1.5ml eppendorf tube.

(Keep a few micoliters to run on a gel against the product, as a negative control)

  • Add water if you need (See example below)
  • Add 10X reaction buffer

(See the enzyme(s) page in the NEB manual or website for the correct one)

  • Add 100X BSA if needed

(BSA is recommended for some reactions (see manual), and will not hurt any)

  • Add restriction enzyme(s) last

(Keep these on ice or in a freezer box, Always) (Also, keep the percentage of enzyme in the reaction below 5% by volume to avoid nonspecific cutting)

  • Let the reaction run at the temperature recommended in the NEB manual for at least 2 hours.

(Overnight is ok)

  • Run a bit of the digest sample against the undigested control to confirm that the digstion worked.

Sample single digest (50 ul total volume)

  1. 30ul DNA
  2. 13.5ul Water
  3. 5ul 10X Buffer
  4. 0.5ul 100X BSA
  5. 1ul Restriction Enzyme

Sample double digest (50 ul total volume)

First, check the NEB manual double digest page for optimal conditions or whether it is recommended against for your enzymes

  1. 30ul DNA
  2. 12.5ul Water
  3. 5ul 10X Buffer (Check NEB [http://www.neb.com/nebecomm/DoubleDigestCalculator.asp double digest table] - many not be what is used for the single digests
  4. 0.5ul 100X BSA (Add if it is required for either enzyme)
  5. 1ul Restriction Enzyme 1
  6. 1ul Restriction Enzyme 2

PCR protocol

Colony PCR with Platinum Taq

  1. Touch the tip of a p2 pipet tip gently to the center of one colony
  2. Swish the tip around in 100uL of sterile water
  3. Vortex for a few seconds then hold on ice
  4. Thaw PCR SuperMIx and keep it on ice
  5. Reconstitute all primers to 100μM
  6. Make a 1:5 working dilution of your primer from the 100μM stock, using sterile water

PCR reactions set up

Reaction PCR SuperMix (uL) Template from colony (uL) 5' Primer (uL) 3' Primer (uL) Water to 50 uL
A 451112
B 45311--
  1. Set up an A and a B reaction for each colony and for each primer set (ps)
    1. ex: colony #1 1-A-PS1, 1-B-PS1, 1-A-PS2, 1-B-PS2
  2. Cycle:
    1. 94 C for 30 sec
    2. 55 C for 30 sec
    3. 72 C for 1 min/ kb of sequence
  3. Cycle 35 times
  4. hold at 4 C when cycles are complete

PCR with Phusion polymerase

Rx mix:

  • 10μL 5x HF buffer
  • 1μL 10mM dNTPs
  • 1.25μL each primer
  • 0.5μL template
  • 0.5μL Phusion polymerase
  • 35.5μL H2O


Initial denaturation: 30s @ 98°C (2:30 if doing colony PCR)

Denaturations: 10s @ 98°C

Annealing: 30s

Extension: 15-30s/kb @ 72°C (20s is usually enough for plasmids)

Colorimetric Assays

LBB Protocol (from Colleen)

LBB Reagent: 0.04% LBB in 45 mM acetic acid

  • (0.04 g LBB in 100 mL 45 mM acetic acid)
  • 45 mM acetic acid (2.81 mL of concentrated (16M) acetic acid in 100 mL)


STANDARDS

Mn(III/IV) standard made with potassium permanganate. Add 2mM stock concentrations in 25 mL.

Standards (uM) 2 mM Stock (uL)
10 125
15 187.5
20 250
40 500
50 625


For Reaction:

  1. Add standard (or sample) to LBB at 1:3 (0.25 mL standard:0.75 mL LBB)
  2. Use DI water as blank before running standard curve
  3. Incubate in dark for 15 minutes
  4. Record Absorbance on a spec at 620 nm


Note: The linear range for the LBB colorimetric method is 10-50 uM. If your samples are above this limit, dilute accordingly to bring into linear range. Do not force standard curve through zero – there is a background for the LBB solution.


[http://www.genewiz.com/preparesample.aspx Setting up Samples for Sequencing]

BioBrick Standard Assembly

' Prefix Suffix
InsertESXP
VectorEXSP

Protocol for Mutant Strand Synthesis Reaction

PROTOCOL

Notes Ensure that the plasmid DNA template is isolated from a dam+ E. coli strain. The majority of the commonly used E. coli strains are dam+. Plasmid DNA isolated from dam– strains (e.g. JM110 and SCS110) is not suitable.

To maximize temperature-cycling performance, Stratagene strongly recommends using thin-walled tubes, which ensure ideal contact with the temperature cycler’s heat blocks. The following protocols were optimized using thin-walled tubes.

1. Synthesize two complimentary oligonucleotides containing the desired mutation, flanked by unmodified nucleotide sequence. Purify these oligonucleotide "primers" prior to use in the following steps (see Mutagenic Primer Design).

2. Prepare the control reaction as indicated below:

5 μl of 10× reaction buffer (see Preparation of Media and Reagents)

2 μl (10 ng) of pWhitescript 4.5-kb control plasmid (5 ng/μl)

1.25 μl (125 ng) of oligonucleotide control primer #1 [34-mer (100 ng/μl)]

1.25 μl (125 ng) of oligonucleotide control primer #2 [34-mer (100 ng/μl)]

1 μl of dNTP mix

39.5 μl of double-distilled water (ddH2O) to a final volume of 50 μl

Last, add: 1 μl of PfuTurbo DNA polymerase (2.5 U/μl)

3. Prepare the sample reaction(s) as indicated below:

Note Stratagene recommends setting up a series of sample reactions using various concentrations of dsDNA template ranging from 5 to 50 ng (e.g., 5, 10, 20, and 50 ng of dsDNA template) while keeping the primer concentration constant.

5 μl of 10× reaction buffer

X μl (5–50 ng) of dsDNA template

X μl (125 ng) of oligonucleotide primer #1

X μl (125 ng) of oligonucleotide primer #2

1 μl of dNTP mix

ddH2O to a final volume of 50 μl

Last, add: 1 μl of PfuTurbo DNA polymerase (2.5 U/μl)

Testing Inducible Lac System

  1. Grow overnight culture of cells (including positive control w/o repressor and negative control w/o fluorescence)
  2. In morning do a 1:20 dilution in LB w/ resitance markers
  3. Incubate for ~2hrs
  4. OD and measure fluorescence (for GFP ex 489 and em 509) - OD levels should be between .3 and .5
  5. Add 1mM IPTG to half of each sample
  6. Incubate for 2hrs
  7. OD and measure fluorescence

Pouring LB plates w/ antibiotics

  1. Heat LB-agar in microwave until it melts - take it out when it starts to boil (about 4 min). Be careful to make sure it doesn't boil over
  2. Gently swirl media and let cool to below 45-50°C (able to hold w/o pain for 10 s). Try to avoid creating too many bubbles. Adding amp at a higher temperature will ruin it. To speed the cooling process, you can swirl it in ice water; however, if you let the temperature drop too low the agar will begin to harden and you will have to re-melt it.
  3. All following steps must be done by a flame
  4. Once cool enough add antibiotic in a 1:1000 dilution (Amp, Sm, Kan, Cm are all stored as 1000x stock)
  5. Quickly pour into petri dishes- add enough to each plate so that the agar just covers the entire plate
  6. Flame bubbles or move them to the side of plate by gently tilting plate
  7. Leave plates on table until they solidify with lids on
  8. Label sleeve or individual plates. Kan= kanamycin, Amp= ampicillin, Cm= chloramphenicol, Sm= spectomycin

Ethanol precipitation of Takara ligations

  1. Place a 15mL falcon of 200 proof ethanol in ice (there's an aliquot in -20°C most likely)
  2. Put a microcentrifuge into a 4°C fridge (you might have to take the rocker out of the one in the small room)
  3. Add 1/10 volume's (of original ligation) worth of pH 5.2 3M sodium acetate; mix well
  4. Add 2 volume's (of original ligation) worth of ice cold 200 proof ethanol; mix well
  5. Stick in -80°C for 15-20 minutes
  6. Spin at max speed (at least 13,000 rpm) for 15-20 minutes at 4°C
  7. Remove supernatant- be careful not to remove the DNA, which may or may not be a visible pellet at the bottom of the tube. It's better to leave some ethanol than to lose your DNA.
  8. Add 1mL 70% EtOH; mix by vortexing briefly
  9. Spin at max speed in microfuge for 1min
  10. Remove supernatant- be careful not to remove the DNA, which may or may not be a visible pellet at the bottom of the tube. It's better to leave some ethanol than to lose your DNA.
  11. Use the vacuum centrifuge (on widget group's bench) to evaporate the remaining ethanol
  12. Resuspend in 10-15μL sterile nuclease-free water. Be sure to pipet the water repeatedly onto the sides of the bottom of the tube and vortex gently a few times. Spin down everything.
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