Difference between revisions of "Team:Michigan/Experiments"

Experiments

Building the Attack/Guard Assay:

SaCas9, gRNA for SaCas9, and gRNA for SpCas9 were synthesized by Integrated DNA Technologies. SaCas9 was synthesized from two linearized fragments, with a Ava restriction site near the 5’ end of one and 3’ end of the other to join them into one fragment. This complete fragment was flanked by EcoRI and PstI sites in order to be compatible for insertion into the submission vector pSB1C3. Once the SaCas9_C plasmid was confirmed, we double digested the SaCas9_C and the gRNA for SaCas9 with SacI and SgrAI and then ligated the fragments, leaving us with one of our final plasmids needed for experiments, SaCas9_C_gRNA, a plasmid containing a LacI coding region for inducible control, the gRNA able to transcribe and bind with SaCas9 in order to target the complementary sequence on the reporter plasmid, and the SaCas9 coding region.


We obtained the SpCas9 DNA from the Zhang lab at the University of Michigan in a pET083 backbone. In order to move the fragment we needed (containing LacI repressor and SpCas9) into pSB1K3, we used PCR mutagenesis to flank our region of interest with EcoRI and PstI cut sites. Once SpCas9_K was confirmed, we double digested with the gRNA for SpCas9 using sites EcoNI and SgrAI. Then we ligated the resulting fragments, leaving us with our final plasmid needed for experiments, SpCas9_K_gRNA, a plasmid containing a LacI Repressor coding region for inducible control, the gRNA able to transcribe and bind with SpCas9 in order to target the complementary sequence on the reporter plasmid, and the SpCas9 coding region.

Given more time to conduct experiments, we would induce the necessary point mutations in each Cas9 through the use of One-Pot Mutagenesis (see protocol section). This requires primers be designed to anneal to the specific portion of the the sequence where we want to induce our mutations. These mutations will be specific to each Cas9 used since the nuclease domain will be in a different location within the sequence in each case.

The reporter plasmid (with CFP) was synthesized by Integrated DNA Technologies to contain the necessary target sequence and PAM sequence needed for recognition of SpCas9 and SaCas9 with their respective gRNAs.

All restriction digests were performed using New England Biolabs (NEB) restriction enzymes. Qiagen PCR cleanup kits were then used on each restriction digest product. All ligations were performed with NEB T4 ligase at room temperature overnight, then the ligation products were first transformed into homemade DH5a chemically competent cells (see protocol section), but after a couple failed transformations, we used NEB DH5a chemically competent E. coli cells. These transformations were plated and grown overnight. 5mL liquid cultures were prepared from colonies picked from the plates the next day and themselves grown overnight. All DNA extraction was performed using Qiagen miniprep kits. CFP was measured at 398 nm excitation and 498 nm emission via a plate reader (Infinite 200, Techan Life Sciences). Cell density was measured at A600 via the same plate reader.

Protocols

Experiment 1:

Will SpCas9 + Reporter and SaCas9 + Reporter effectively degrade the Reporter plasmid, reducing CFP over time?
Setup:
Start with 3 types of 5mL liquid cultures grown overnight shaking 225 rpm at 37C: Reporter plasmid (CFP - Amp), SpCas9 with gRNA and Reporter with target sequence (Kan + Amp), and SaCas9 with gRNA and Reporter with target sequence (Chlor + Amp).

1. Add 4mL prewarmed (37C) LB-Amp (2 15mL confocal tubes), add 4mL prewarmed (37C) LB-Kan + Amp (3 15mL confocal tubes), and add 4mL prewarmed (37C) LB-Chlor + Amp (3 15mL confocal tubes)
2. Add 1mL from each 5mL starter culture to three 4 mL of prewarmed media for SpCas9 and SaCas9 (plus the Reporter CFP plasmid for each) and two 4 mL of prewarmed media for the Reporter CFP plasmid.
3. Grow all 8 5mL cultures at 37C until A600 is between 1.0 and 1.2.
4. When all cultures are between 1.0 and 1.2, record A600 for each culture (Time 0), measure CFP and image fluorescence.
5. Add IPTG to a final concentration of 1 mM to two cultures of SpCas9 and SaCas9 and to one tube of the Reporter plasmid.
6. Every 15 minutes take measurements of A540 for all cultures, measure CFP (via plate reader), and image fluorescence for 1 hour.
7. Every 30 minutes take measurements of A540 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.
8. Every 1 hour take measurements of A540 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.

Experiment 2:

Will dSpCas9 + Reporter and dSaCas9 + Reporter result in little to no changes in CFP level compared to control?
Setup:

Start with 3 types of 5mL liquid cultures grown overnight shaking 225 rpm at 37C: Reporter plasmid (CFP - Amp), dSpCas9 with gRNA and Reporter with target sequence (Kan + Amp), and dSaCas9 with gRNA and Reporter with target sequence (Chlor + Amp).
1. Add 4mL prewarmed (37C) LB-Amp (2 15mL confocal tubes), add 4mL prewarmed (37C) LB-Kan + Amp (3 15mL confocal tubes), and add 4mL prewarmed (37C) LB-Chlor + Amp (3 15mL confocal tubes)
2. Add 1mL from each 5mL starter culture to three 4 mL of prewarmed media for dSpCas9 and dSaCas9 (plus the Reporter CFP plasmid for each) and two 4 mL of prewarmed media for the Reporter CFP plasmid.
3. Grow all 8 5mL cultures at 37C until A600 is between 1.0 and 1.2.
4. When all cultures are between 1.0 and 1.2, record A600 for each culture (Time 0), measure CFP and image fluorescence.
5. Add IPTG to a final concentration of 1 mM to two cultures of dSpCas9 and dSaCas9 and to one tube of the Reporter plasmid.
6. Every 15 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 1 hour.
7. Every 30 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.
8. Every 1 hour take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.

Experiment 3:

What are the results with E.Coli that have been transformed with SpCas9, dSacas9, and Reporter?
Setup:
Start with 2 types of 5mL liquid cultures grown overnight shaking 225 rpm at 37C: Reporter plasmid (CFP - Amp), SpCas9 with gRNA, Reporter with target sequence (Kan + Amp), and dSaCas9 with gRNA (Chlor + Amp + Kan).
1. Add 4mL prewarmed (37C) LB-Amp (2 15mL confocal tubes) and add 4mL prewarmed (37C) LB-Chlor + Amp + Kan (4 15mL confocal tubes)
2. Add 1mL from each 5mL starter culture to four 4 mL of prewarmed media for E.coli containing SpCas9, dSaCas9, and reporter plasmids and two 4 mL of prewarmed media for the Reporter CFP plasmid.
3. Grow all 6 5mL cultures at 37C until A600 is between 1.0 and 1.2.
4. When all cultures are between 1.0 and 1.2, record A600 for each culture (Time 0), measure CFP and image fluorescence.
5. Add IPTG to a final concentration of 1 mM to two cultures of the E.coli containing SpCas9, dSaCas9, and reporter plasmids and to one tube of the Reporter plasmid.
6. Every 15 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 1 hour.
7. Every 30 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.
8. Every 1 hour take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.

Experiment 4:

What are the results with E.Coli that have been transformed with SaCas9, dSpCas9, and Reporter?
Setup:
Start with 2 types of 5mL liquid cultures grown overnight shaking 225 rpm at 37C: Reporter plasmid (CFP - Amp), dSpCas9 with gRNA, Reporter with target sequence (Kan + Amp), and SaCas9 with gRNA (Chlor + Amp + Kan).
1. Add 4mL prewarmed (37C) LB-Amp (2 15mL confocal tubes) and add 4mL prewarmed (37C) LB-Chlor + Amp + Kan (4 15mL confocal tubes)
2. Add 1mL from each 5mL starter culture to four 4 mL of prewarmed media for E.coli containing dSpCas9, SaCas9, and reporter plasmids and two 4 mL of prewarmed media for the Reporter CFP plasmid.
3. Grow all 6 5mL cultures at 37C until A600 is between 1.0 and 1.2.
4. When all cultures are between 1.0 and 1.2, record A600 for each culture (Time 0), measure CFP and image fluorescence.
5. Add IPTG to a final concentration of 1 mM to two cultures of the E.coli containing dSpCas9, SaCas9, and reporter plasmids and to one tube of the Reporter plasmid.
6. Every 15 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 1 hour.
7. Every 30 minutes take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.
8. Every 1 hour take measurements of A600 for all cultures, measure CFP (via plate reader), and image fluorescence for 2 hours.

Agarose Gel Electrophoresis:

1. Fill an erlenmeyer flask with 50ml of 1X TAE buffer.
2. Weigh out and add 0.75g of agarose to make 1.5% gel. Mix by swirling the flask
3. Microwave for ~2.5 minutes without boiling. Then run the beaker under cold water until the glass has cooled.
4. Add 5uL of GelRed and mix gently by swirling.
5. Pour the beaker’s contents into the gel mold.
6. Add plastic “combs” to the mold to make wells.
7. Push any large bubbles to the side walls if possible using a spare comb or pipet tip.
8. Let the gel cool and solidify (~10-20 min).
9. Once the gel is solid, move it into the electrophoresis device. Make sure the ‘top’ end of the gel (with the wells) is on the same side as the negative terminal.
10. Pour 1X TAE buffer to the fill line.
11. Slowly and gently remove the plastic combs from the wells.
12. Select a ladder to use based on what kind of samples you’re running.
13. Load 10uL of the ladder into the leftmost well.
14. For each sample, in a small PCR tube, add 10uL of the DNA sample and 2uL of 6X purple loading dye and mix by pipetting up and down.
15. Load each sample into a well in the same way you loaded the ladder.
16. Run the gel.

Transforming DH5-Alpha strain E. coli Cells:

1. Warm the water bath to 42C and set shaker to 37C.
2. Thaw the plasmid DNA on ice.
3. Thaw one 50 uL vial of DH5-Alpha cells on ice.
4. Pipet 1-5 uL of the plasmid into the vial of cells, mix by gently tapping side of tube.
5. Store the extra plasmid at -20 C.
6. Put the tube of cells and DNA in ice for 30 minutes.
7. Make sure the hot water bath is at 42 C.
8. Put the vial in the 42 C water bath for exactly 30 seconds. Do not mix or shake.
9. Immediately put the vial back on ice for 5 min.
10. Add 950 uL of room temperature SOC medium to the vial.
11. Tape the tube of cells and DNA to a shaking incubator, and shake/incubate at 37 C and 225 rpm for exactly an hour.
12. Transfer 10 uL and 500 uL of the culture onto two petri dishes with correct antibiotic for the plasmid.
13. Spread evenly on plates using disposable sterile spreader.
14. Incubate the plates overnight (~12-20 hrs) at 37 C.

Miniprep:

We use Quiaprep spin kits for our minipreps.
1. Transfer 1.5 ml of the bacterial culture to a 1.5 ml eppendorf tube and centrifuge for 5 min at 6,000 rpm. Dump out supernatant from the eppendorf tube and add an additional 1.5 ml of culture. Repeat until your whole culture has been pelleted.
2. Pour out the supernatant into a liquid waste container.
3. Add 250uL of Buffer P1 to the tube containing the pellet, then pipet up and down until the pellet is completely resuspended. When you can no longer see any trace of the pellet, expel all liquid from the pipet back into the tube and proceed.
4. Add 250uL of Buffer P2.
5. With the lid closed flip the tube 4-6 times to mix. Wait 1 minute before proceeding.
6. Add 350uL of Buffer N3, then mix again, flipping it 4-6 times.
7. Centrifuge for 10 minutes at 13,000 rpm.
8. The kit comes with its own blue open top tubes with filters in them. Get one of these. There are two parts, the inner part with a filter and the outer part that’s just a tube. Label both of these parts to avoid any mix ups.
9. Transfer as much of the supernatant as you can (usually between 800uL and 1000uL) from the tube you just centrifuged onto the filter of the tube from the kit
10. Centrifuge the new tube with the filter in it for 60 seconds at 13,000 rpm.
11. Detach the bottom/outer part of the tube, dump its contents but keep the tube itself.
12. Put the inner part with the filter back inside the now empty outer part. Drip 500uL of Buffer PB onto the filter.
13. Centrifuge for 60 seconds at 13,000 rpm.
14. Detach the outer part and discard the flow-through again, then put the inner part back inside.
15. Drip 750uL of Buffer PE into the filter, again taking care not to touch the filter.
16. Centrifuge for 60 seconds. Discard the flow-through.
17. Centrifuge again for 60 seconds.
18. Discard the whole outer part of the tube, placing the inner part in a new, clean, and labeled eppendorf tube.
19. Drip 50uL of Buffer EB onto the filter, again taking care not to touch the filter.
20. Let it stand for 1-2 minutes.
21. Centrifuge for 60 seconds.
22. Throw out the inner portion of the tube.
23. Store the DNA in the freezer.

Restriction digest:

1. In a PCR tube, add 1ug of the DNA you want to digest.
2. Add 5uL of the 10X NEB Buffer you selected to match the restriction enzyme you’re using.
3. Add an amount of ddH2O which will give you 50ul final volume for your reaction.
4. Add 1ul of your chosen restriction enzyme. Stir the liquid with the pipette tip to make sure everything is fully mixed.
5. Incubate for 1 hour at the temperature specified by the restriction enzyme you’re using.
6. If the restriction enzyme you’re using is heat innactivatable then heat your reaction at the recommended temperature for 20 minutes.
7. If your enzyme is not heat innactivatable, you must use a PCR cleanup kit to remove your restriction enzyme.

Ligation:

1. Add 2uL of T4 DNA Ligase Buffer to a microcentrifuge tube.
2. Add 50ng of the vector DNA to the tube.
3. Add 37.5ng of the insert DNA to the tube.
4. Add nuclease free water to the tube so that the total volume is 19uL.
5. Add 1uL of T4 DNA Ligase. Gently mix the reaction by stirring with the pipette tip.
6. Incubate at 16C overnight or 1 hr at room temperature.
7. Put in 65C hot water bath for 10 minutes.
8. Chill on ice, then either transform into cells or store at -20C.

PCR:

1.Determine the number of PCR reactions you will be performing, then add 2.
2.Multiply all the following amounts by this new number to create the “master mix” for the PCR. Round up to the nearest uL. In a microcentrifuge tube add:
a) 25uL Q5 2X high fidelity master mix (NEB)
b)0.25uL 100uM forward primer
c)0.25uL 100uM reverse primer
d) 23.75uL Nuclease Free Water
3. Mix the master mix by pipetting up and down gently.
4. In small PCR tubes, on ice, add 2 uL (from 1-10 ng/ul stock for a 5kb plasmid) of the template DNA from each sample.
5. Add 48uL of the master mix to each PCR tube.
6. Load the PCR tubes onto the heat cycler.
7. Heat cycle through the program designed for your primers.

Gibson Assembly

1. In a PCR tube, add the following:
a) Xul of each of your gibson compatible fragments
b) 10uL 2X gibson assembly master mix
c) ddH2O so that the total volume is 20uL
2. In a second PCR tube, add the following:
a) 10ul of Positive Control mix
b) 10uL 2X gibson assembly master mix
3. Incubate the two tubes in the thermocycler at 50C for 15 minutes, then either store in the -20C freezer or immediately transform into NEB 5-alpha cells

Gel Imaging

1. Put blue plastic test plate on the imaging tray. Put the gel on the middle of the black part.
2. Move camera cone and cover the imaging tray. Cover completely.
3. Turn on the UV bulb.
4. Touch the “live” button on the computer screen. You should see the fake gel lines glow.
5. Turn bulb off.
6. Remove camera cone, replace the fake gel with your real gel.
7. Replace the camera cone.
8. Turn bulb on.
9. Keep adjusting camera until you can see the gel clearly.
10. Touch the “snap” button to capture your image.
11. Touch the “save button.”
12. Turn the bulb off, turn the imager off, wipe down the imaging tray and toss your gel.

Making LB Media

1. Get a glass screwtop container about twice the size of the amount of media you want to make.
2. Add the same amount of deionized water as you want media
3. Weigh out 25g of LB broth powder per 1L of media you want to make and add it to the flask.

• 2.5g for 100mL
• 5g for 200mL
• 6.25g for 250mL
• 12.5g for 500mL

4. Swirl/vortex until mostly dissolved
5. Partially screw on the top and put a piece of autoclave tape on it.
6. Autoclave on appropriate liquid setting for the amount of liquid you have.
7. Let cool until at room temperature
8. If you are using the media immediately, you must wait until it has cooled to a temperature where you can comfortably touch the container before adding antibiotic.
9. Store at room temp with screw top completely closed.

Making LB Plates

1. Each plate takes 20-25mL of media.
2. Get a glass flask about twice the size of the amount of media you want to make.
3. Add the same amount of deionized water as you want media.
4. Weigh out 32g of LB agar powder per 1L of agar you want to make and add it to the flask.

• 3.2g for 100mL
• 8g for 250mL
• 16g for 500mL

5. Swirl/vortex flask until mostly dissolved.
6. Cover top in tin foil and put a piece of autoclave tape on it.
7. Autoclave on liquid setting.
8. Let cool until you can touch glass.
9. Add proper amount of antibiotic.
10. Pour evenly into plates. Once you pour into a plate, cover it ¾ of the way with its lid.
11. Let the plates solidify. Once they are solid, put the lids on all the way, and store them upside down in the fridge.

NEB Gel Extraction

1. Use a razor blade to cut the band you want out of the gel. Minimize the exposure of the gel to UV light since extra exposure damages the DNA.
2. Weigh the gel slice, then put the gel slice into a microcentrifuge tube.
3. Add 4uL of the Gel Dissolving Buffer per 1mg of the gel.
4. Incubate the tube in a 50C water bath, vortexing it occasionally until the gel is completely dissolved.
5. Get an open top tube with a filter in it.
6. Put the filter inside the tube, the add the now dissolved gel solution to the filter.
7. Spin in the microcentrifuge at 13,000 rpm for 1 minute.
8. Detach the filter part and discard the contents of the outer part of the tube.
9. Put the filter part back in the outer tube, then add 200uL DNA Wash Buffer.
10. Spin for another minute at 13,000 rpm.
11. Discard the contents of the outer tube again.
12. Spin again for a minute at 13,000 rpm.
13. Discard the contents of the outer tube again.
14. Take the inner filter part out and put it in a new, clean, microcentrifuge tube.
15. Label the new tube with as much detail as possible, this is the final storage tube for your DNA.
16. Add 10uL of Elution Buffer to the filter.
17. Wait one minute.
18. Spin at 13,000 rpm for a minute.
19. Take the filter out and discard it. The liquid in the microcentrifuge tube is your purified DNA.

Making E.coli competent cells

1. Inoculate a single colony into 5mL Lb in 50mL falcon tube. Grow O/N @ 37°C.

 2. Use 1mL to inoculate 100mL of LB in 250mL bottle the next morning. 3. Shake @ 37°C for 1.5-3hrs.


 Or

3. Inoculate a single colony into 25mL LB in a 250 mL bottle in the morning.

 4. Shake @ 37°C for 4-6 hrs.


 Then….

5. Put the cells on ice for 10 mins (keep cold form now on).

 6. Collect the cells by centrifugation in the big centrifugue for 3 mins @6krpm

 7. Decant supernatant and gently resuspend on 10 mL cold 0.1M CaCl (cells are susceptible to mechanical disruption, so treat them nicely).

 8. Incubate on ice x 20 mins

 9. Centrifuge as in 2

 10. Discard supernatant and gently resuspend on 5mL cold

 11. 0.1M CaCl/15%Glycerol

 12. Dispense in microtubes (300μL/tube). Freeze in -80°C.

One-Mutagenesis

Wrenbeck, E., Klesmith, J., Stapleton, J., & Whitehead, T. (2016). Nicking mutagenesis: Comprehensive single-site saturation mutagenesis.

Kinase mutagenic oligos and secondary primer

1. Make a mixture of NNN/NNK mutagenic oligos at a final concentration of 10 μM.
2. Into a PCR tube, add:
20 μL 10 μM mutagenic oligo mixture
2.4 μL T4 Polynucleotide Kinase Buffer
1 μL 10 mM ATP
1 μL T4 Polynucleotide Kinase (10 U/μL)
3. In a separate PCR tube add:
18 μL NFH2O
3 μL T4 Polynucleotide Kinase Buffer
7 μL 100 μM secondary primer
1 μL 10 mM ATP
1 μL T4 Polynucleotide Kinase (10 U/μL)
4. Incubate at 37°C for 1 hour. Store phosphorylated oligos at -20°C. The day of mutagenesis, dilute phosphorylated mutagenic oligos 1:1000 and secondary primer 1:20 in NFH2O.

Prepare ssDNA template

5. Add the following into PCR tube(s):
0.76 pmol plasmid dsDNA
2 μL 10X CutSmart Buffer
1 μL 1:10 diluted Exonuclease III (final concentration of 10 U/μL)
1 μL Nt.BbvCI (10 U/μL)
1 μL Exonuclease I (20 U/μL)
NFH2O to 20 μL final volume
6. Run the following PCR program:
37°C 60 minutes
80°C 80 minutes
4-10°C Hold

Single-site saturation mutagenesis strand 1

7. Add the following into each tube (100 μL final volume):
26.7 μL NFH2O
20 μL 5X Phusion HF Buffer
4.3 μL 1:1000 diluted phosphorylated mutagenic oligos
20 μL 50 mM DTT
1 μL 50 mM NAD+
2 μL 10 mM dNTPs
1 μL Phusion High Fidelity Polymerase (2 U/μL)
5 μL Taq DNA Ligase (40 U/μL)
8. Run the following PCR program:
98°C 2 minutes
————————-
15 cycles of:
98°C 30 seconds
55°C 45 seconds
72°C 7 minutes
***add additional 4.3 μL oligo at beginning of cycles 6 and 11***
————————-
45°C 20 minutes
4-10°C Hold

Column purification I

9. Following the manufacturers’ instructions, perform a column purification using a Zymo Clean and Concentrate Kit:
a) Add 5 volumes of DNA binding buffer to each reaction and mix
b) Transfer to a Zymo-Spin Column in a collection tube
c) Centrifuge at maximum speed for 30 seconds and discard flow through
d) Add 200 μL of DNA wash buffer to the column
e) Centrifuge at maximum speed for 30 seconds and discard flow through
f) Repeat steps 4 and 5
g) Add 15 μL of NFH2O directly to the column in a new clean 1.5 mL microfuge tube and incubate at room temperature for 5 minutes
h) Centrifuge at maximum speed for one minute

Degrade template strand

10. Transfer 14 μL of the purified DNA product to a PCR tube, then add (20 μL final volume):
2 μL 10X CutSmart Buffer
2 μL 1:50 diluted Exonuclease III (final concentration of 2 U/μL)
1 μL 1:10 Nb.BbvCI (final concentration of 1 U/μL)
1 μL Exonuclease I (20 U/μL)
11. Run the following PCR program:
37°C 60 minutes
80°C 20 minutes
4-10°C Hold

Synthesize 2nd (complementary) mutagenic strand

12. To the above PCR tubes, add (100 μL final volume):
27.7 μL NFH2O
20 μL 5X Phusion HF Buffer
3.3 μL 1:20 diluted phosphorylated secondary primer
20 μL 50 mM DTT
1 μL 50 mM NAD+
2 μL 10 mM dNTPs
1 μL Phusion High Fidelity Polymerase (2 U/μL)
5 μL Taq DNA Ligase (40 U/μL)
13. Run the following PCR program:
98°C 30 seconds
55°C 45 seconds
72°C 10 minutes
45°C 20 minutes
4-10°C Hold

DNA clean up

14. Add into each reaction 2 μL of DpnI (20 U/μL) and run the following PCR program:
60°C 60 minutes

Column purification II

15. Follow the instructions in step 9 but elute in 6 μL of NFH2O.

DNA Transformation

16. Transform the entire 6 μL reaction product into a high-efficiency cloning strain following standard transformation protocols. After recovery, bring the final volume of the transformation to 2-2.5 mL with additional sterile media. Spread on to a prepared large BioAssay dish (245 mm x 245 mm x 25 mm, Sigma-Aldrich). Additionally, serial dilution plates should be prepared to calculate transformation efficiencies. Incubate overnight at 37°C. The next day, scrape the plate using 5-10 mL of LB or TB. Vortex the cell suspension and extract the library plasmid dsDNA using a mini-prep kit (Qiagen) of a 1 mL aliquot of the cell suspension. Additional mini-preps (or a midi-prep) can be done if large amounts of library DNA are required.

See our lab notebook here.