Team:UC Berkeley/Protocols

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Contents

PCR

PCR (25ul)

  1. Mastermix: 20.375 ul water, 0.5ul 10mM dNTP, 2.5 ul 10x Buffer 2, 0.375 ul Expand Polymerase 1. For reactions with DMSO, use 2.5uL of DMSO and 17.875uL of water.
  2. Aliquot 23.75 ul into each tube.
  3. Add 0.5ul Oligo 1 (10uM), 0.5ul Oligo 2 (10uM), 0.25 ul template DNA.
  4. Load into thermocycler. Note: small holes are for 0.2 ul tube and big holes for 0.5 ul to maximize contact
  5. Select the program (55/45 if under 1kb, 2K55/2K45 for 1kb to 2kb, 4K55/45 for 2kb to 4kb, 8K55/45 if over 4kb)

Wobble PCR (50ul)

  1. Mastermix: 40 ul water, 1.5 ul MgCl2 (50mM), 5 ul 10x buffer (Taq), 1 ul 10mM dNTP, 0.5 ul Taq Polymerase
  2. Aliquot 48 ul into each tube
  3. Add 1 ul 100 uM Oligo 1, 1 ul 100 uM Oligo 2.
  4. Load into thermocycler.
  5. Select the program WOBBLE55 or WOBBLE45 and run.

Colony PCR (20ul)

  1. Mastermix: 16.1 ul water, 0.6 ul MgCl2 (50mM), 2 ul 10x buffer (Taq), 0.4 ul 10mM dNTP, 0.1 ul Taq Polymerase, 0.4 ul ca998 oligo (10uM), 0.4 ul g00101 oligo (10uM)
  2. Aliquot 20 ul into each tube
  3. Pick the desired colony with a 10 ul pipette tip and swirl around in the PCR mix before putting into culture tube with appropriate antibiotic.
  4. Load into thermocycler.
  5. Select the program COLONY and run.

Purification and Visualization of PCR Product

  1. Label new tubes with the names of your samples for isolation.
  2. Open a 0.8% CloneWell E-Gel. Only do this unless you are planning on running it in less than 15 min or it will dry out!
  3. Place E-gel in apparatus (don't remove combs yet!) and do a PreRun for 2 minutes.
  4. Remove combs and add about 25 ul of water to all cloning wells, 10 ul of ladder (GeneRuler, 1kb DNA Ladder) to the M lane, and 25 ul of your PCR products to each well. If you have fewer than 8 samples, try not to use the two outer lanes (1 and 8) since they tend to dry out/distort the DNA, and the water in these clone wells evaporated extremely quickly. Also, fill any unused lanes with 25 ul of water.
  5. Select Run CloneWell and hit Go. After about 1 minute, turn on the blue light and view the gel through the orange shield; you should see small, bright bands under your sample lanes. Every couple minutes, check to make sure the clone wells are still filled with water.
  6. Allow the bands to run until they reach the tiny horizontal scratches just above the clone wells. At this point, hit Go to stop the gel from running. Completely fill all clone wells with water and prepare for extraction.
  7. Hit Go to start again and watch until the band has mostly entered the clone well and water. Hit Go again to stop the gel.
  8. Extract the water/sample from the clone well with a pipette and transfer to appropriately-labeled new tube. If you miss your sample and it has continued on past your clone well, you can stop the program and select a reverse run to get it back, but avoid doing this if you can
  9. Go ahead and freeze your samples if you don't plan on doing a digestion immediately.


Digestion (w/o DpnI)

  1. Combine 5 ul NEB2, x ul of your DNA samples from the CloneWell, 1 ul EcoRI, 1 ul BamHI, (43-x) ul water. It's usually safe to assume you'll have at least 15 ul of any given sample extracted from the E-gel, so if you have many samples make a master mix in which there are 15 ul of DNA sample and 28 ul of water and multiply accordingly depending on the number of samples you have.
  2. Incubate at 37 C for one hour.


Clean-up with Zymo Columns

Products Greater Than 300 bp

  1. Label each Zymo column and fresh Eppendorf tubes with the appropriate names of your samples.
  2. Add 200ul ADB buffer to each of the digestion samples, mix and pipette into zymo columns. Spin for 30 sec at full speed to pass the liquid through the column.
  3. Empty collection tubes and put back onto the columns. Wash with 200 ul wash buffer and again spin for 30 sec at full speed. This will dissolve any extra guanidinium chloride and salts sitting on the membrane
  4. Repeat the wash buffer step 3. Now nothing is present on the membrane but the DNA and a little ethanol and water
  5. Now spin the column for 90 sec at full speed to remove all traces of water/ethanol.
  6. Empty and discard the collection tube. Replace each collection tube with the appropriately-labeled fresh Eppendorf tubes.
  7. Add 6 ul of water directly to each of the membranes of the Zymo columns, and spin for 45 sec to elute the DNA into your fresh Eppendorf tubes.

Products Between 20 and 300 bp

  1. Add 1 volume equivalent of Zymo ADB buffer to each reaction. Vortex to mix.
  2. Add 5 volumes of 95% ethanol (under fume hood, labeled flammable). Vortex to mix. The remaining steps are just like a normal Zymo clean-up reaction
  3. Label each Zymo column and fresh Eppendorf tubes with the appropriate names of your samples.
  4. Transfer your reaction mixtures into Zymo columns. Spin for 30 sec at full speed to pass the liquid through the column.
  5. Empty collection tubes and put back onto the columns. Wash with 200 ul wash buffer and again spin for 30 sec at full speed. This will dissolve any extra guanidinium chloride and salts sitting on the membrane
  6. Repeat the wash buffer step 3. Now nothing is present on the membrane but the DNA and a little ethanol and water
  7. Now spin the column for 90 sec at full speed to remove all traces of water/ethanol.
  8. Empty and discard the collection tube. Replace each collection tube with the appropriately-labeled fresh Eppendorf tubes.
  9. Add 6 ul of water directly to each of the membranes of the Zymo columns, and spin for 45 sec to elute the DNA into your fresh Eppendorf tubes.


Ligation

  1. Mastermix: 6.5 ul water, 1 ul ligation buffer, 0.5 T4 DNA ligase, 1 ul pBca1256 digested with EcoRI and BamHI.
  2. Aliquot 9 ul into fresh tubes.
  3. Add 1 ul of your insert.
  4. Cover with foil and incubate for 30 min at room temperature.


Transformation

Remember to pre-warm your plates at 37 degrees C!!

Transformation from Ligation Reaction

  1. Retrieve your competent cells from the -80 (on the third shelf) and thaw on ice. There are 220 ul of competent cells in each tube
  2. Add 30 ul cold KCM and 20 ul cold water to each tube of competent cells. Invert ~2x to mix.
  3. Take the top of an empty pipette tip box, add ice and water. Cut out the appropriate amount of tubes from the sets of attached tubes and float them in this ice bath. Do NOT let any water get into the tubes or else you will have to start over due to possible contamination!
  4. Combine 45 ul of your cells into 10 ul of your ligation rxn while remembering to keep all tubes on ice. Swirl and pipette up and down once to mix.
  5. Foil all tubes and incubate 10 min in ice-water.
  6. Heat shock your cells by placing them in a 42 C water bath for 90 sec.
  7. Remove and incubate on ice for 2 min.
  8. Rub ethanol/flame top of foil to sterilize. Add 50 ul of LB 2YT to each tube by poking holes through the foil with your pipette tips.
  9. Re-cover all tubes with foil and incubate at 37 C for 1 hour.
  10. Plate using the sterilized glass-bead method or with standard spreaders.

Transformation from Mini-prep

  1. Retrieve your competent cells from the -80 (on the third shelf) and thaw on ice. There are 220 ul of competent cells in each tube
  2. Add 30 ul cold KCM and 50 ul cold water to each tube of competent cells. Invert ~2x to mix. This will give you a total of 300 ul of competent cells
  3. Take the top of an empty pipette tip box, add ice and water. Cut out the appropriate amount of tubes from the sets of attached tubes and float them in this ice bath. Do NOT let any water get into the tubes or else you will have to start over due to possible contamination!
  4. Combine 30 ul of your cells into 1 ul of your plasmid prep while remembering to keep all tubes on ice. Swirl and pipette up and down once to mix.
  5. Foil all tubes and incubate 10 min in ice-water.
  6. Heat shock your cells by placing them in a 42 C water bath for 90 sec.
  7. Remove and incubate on ice for 2 min.
  8. Rub ethanol/flame top of foil to sterilize. Add 50 ul of LB 2YT to each tube by poking holes through the foil with your pipette tips.
  9. Re-cover all tubes with foil and incubate at 37 C for 1 hour.
  10. Plate using the sterilized glass-bead method or with standard spreaders.

Transformation into Cell Culture

  1. Grow cells in ~4 mL LB until cloudy (OD600=0.5)
  2. Put on ice
  3. Transfer 1mL into an eppendorf tube on ice, let cool
  4. Centrifuge full speed for 30 sec, toss out supernatant
  5. Resuspend in 90uL of TSS solution
  6. Add 10uL KCM
  7. Add 1uL plasmid DNA
  8. Let sit on ice for 10min, heat shock 90 sec at 42, incubate and/or plate


MiniPrep (1-5 mL)

QIAPrep Spin Miniprep Kit

  1. Pellet 1.5 mL saturated culture by spinning full speed for 30 sec.
  2. Dump supernatant, repeat to pellet another 1.5 mL (for a total of 3 mL)
  3. Add 250 ul of P1 buffer into each tube. Vortex well to resuspend cells. You should not see anything solid at the bottom of the tube
  4. Add 250 ul of P2 buffer (a base that denature everything and causes cells to lyse). Gently invert until uniformly light blue.
  5. Add 350 ul of N3 buffer (an acid of pH ~5 that cuases cell junk, including protein and chromosomal DNA, to precipitate and leaves plasmids and other small molecules in solution). Slowly invert ten times.
  6. Spin in centrifuge at top speed for 5 min.
  7. Label blue columns with an alcohol-resistant lab pen.
  8. Pour liquid into columns, and place the columns into the centrifuge. Spin at max speed for 30 sec.
  9. Dump liquid out of the collection tubes into waste container. The DNA should be stuck to the white resin now
  10. Wash each column with 500 ul of PB buffer.
  11. Spin in centrifuge at max speed for approximately 15 sec, then flick out the liquid again.
  12. Wash with 750 ul of PE buffer. This washes the salts off the resins
  13. Spin in centrifuge at max speed for approximately 15 sec, then flick out the liquid again.
  14. Spin one more time at max speed for 90 sec to dry out resin.
  15. Label new tubes and put columns in them. Discard collection tubes.
  16. Elute them by squirting 50 ul of water directly onto the resin in each column. Get as close to the resin with your pipette tip as possible without touching it
  17. Spin in centrifuge at top speed for 30 sec.
  18. Take out columns and cap the tubes. Check to make sure there is something in the tube! It's hard to keep track of which tubes you added water to when working with several samples. If there is nothing in the tube, just add your water and spin again.
  19. Clean-up. Note: the P1 buffer is stored at 4 C and all the rest are stored at room temp

Agencourt Miniprep

  1. Pellet 800 ul 2YT saturated cultures
  2. Resuspend in 100 ul RE1
  3. Add 100 ul L2 to alkaline lyse
  4. Add 100 ul N3
  5. Wait 10 min for precipitate to float out, or spit it out
  6. Transfer 110 ul of lysate to round bottom microtiter plate
  7. Add 10 ul of PUR4 beads
  8. Add 80 ul of isopropanol
  9. Pipette 10x to mix
  10. Allow plate to separate on magnet for 15 min
  11. Dump supernatant
  12. Add 200 ul of 70% ethanol while still on magnet
  13. Let incubate for 30 sec, then discard
  14. Do the above 70% ethanol wash 3x
  15. Let beads dry for 30 min
  16. Add 40 ul RE1 and shake to elute
  17. Aspirate over liquid

Transfer of Basic Parts into Assembly Vectors Using Eco/Bam

Digesting Basic Part in Entry Vector

Set up the digest - make a master mix of 21 uL water, 3 uL NEB2 buffer, .5 uL EcoRI, and .5 uL BamHI per reaction. Aliquot 25 uL, then add 5 uL miniprepped plasmid. Incubate for 45 min. at 37 degrees C.

Cleanup the Digest

Remember to make fresh 70% ethanol before starting

Add 70 uL RE1, 5 uL SPRI beads, and 75 uL isopropanol. Pipette up and down 10 times slowly and carefully. Transfer to the magnet plate and wait for 5 minutes. Pipette out the liquid, being careful to stick pipette tips straight down through the ring of magnetic beads. Fill wells with 70% ethanol, wait 30 seconds, pipette out the ethanol. Repeat once. Let beads dry for 30 minutes.

Setting up the Ligation

Make a ligation master mix #1 with 1 uL predigested assembly vector, 1.5 uL ligase buffer, and 12.5 uL water. Add 15 uL to each tube, scraping the beads down with your pipette tip. Then make ligation master mix #2 with .3 uL ligase, .5 uL ligase buffer, and 4.2 uL water. Add 5 uL to each tube, pipetting up and down to mix. Cover with foil, vortex, and incubate at room temperature for 30 minutes.

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