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Assignments Due

• Arnold (Annl Rev Biophys Biomol Struct 2007, 36:1-19)
• Collins (Nat Biotechnol 2006, 24(5):545-554)
• BioBricks

Lecture Topics

• Wiring parts to function collectively: Joff Silberg (Invited Speaker)
• Registry of Standard Biological Parts

Overview of Experiment

Background

• DNA Mini Prep
• Restriction Enzymes

Plasmid mini prep

Disposal of Waste:
• Discard bacterial supernatant in a small beaker; add bleach to 10% for 10 minutes before dumping into the sewer
• Collect contaminated tips and culture tubes in a beaker at your desk then rinse the beaker with bleach after dumping the waste in the clear Biohazards bag; these bags will be autoclaved prior to placement in the household trash
• After the bacteria are lysed, tips, vials, and other materials should be discarded in the regular trash

1. Centrifuge 1.5 ml of bacterial culture for 30 seconds at maximum speed (in a microcentrifuge); discard the supernatant (into small waste beaker)
2. REPEAT step 1.
   
   NOTE: typically you would save a small amount of culture to streak a plate and prepare a fresh overnight for a glycerol stock; bacterial cultures can be stored indefinitely at -80°C without significant loss of viability in media containing at least 15% glycerol (v/v).
   
   
3. Add 600 µl nuclease-free water to the cell pellet and resuspend completely by gently pipetting up and down
4. Add 100 µl of 7X Lysis Buffer (Blue) and mix by inverting tube 6 times
   • Complete lysis is indicated when the solution changes from opaque to clear blue
   • You must proceed to step 5 within 2 minutes (excessive lysis can denature plasmid DNA!)
5. Add 350 µl COLD Neutralization Buffer (Yellow), containing 100 µg/ml RNaseA, and mix thoroughly by inverting tube
   • When neutralization is complete, sample turns yellow and a yellowish precipitate forms
   • Invert sample 2-3 more times (to ensure complete neutralization)
6. Centrifuge at 16,000 x g for 4 minutes
7. Transfer supernatant to a Zymo-Spin™ II column (avoid disturbing pellet!)
8. Put column in a 2 ml collection tube and centrifuge for 15 seconds (maximum speed, press and hold the "short" button)
9. Discard flow-through and put column back into the collection tube
10. Add 200 µl Endo-Wash Buffer to column and centrifuge for 15 seconds (maximum speed, press and hold the "short" button)
11. Add 400 µl Zyppy™ Wash Buffer (containing ethanol) to the column and centrifuge for 30 seconds (maximum speed)
12. Transfer column into a sterile 1.5 ml tube
13. Add 30 µl nuclease-free water directly to the column matrix and wait one minute (room temperature)
14. Centrifuge 15 seconds (maximum speed, press and hold the "short" button) to elute plasmid DNA

Restriction enzyme digests of mini prep DNA

Uncut control (UC), XbaI (20 units/µl) digest (X), PstI (20 units/µl) digest (P), XbaI/PstI double digest (X/P)
1. Put 5 µl plasmid DNA (BBa_I13522) in each tube
2. Add 13 µl nuclease-free water to the double digest; add 14 µl nuclease-free water to the uncut control and single digests
3. Add 2.5 µl 10X NEB buffer #3 to each tube
4. Add 2.5 µl 10X BSA to each tube
5. Add
   • 1 µl nuclease-free water to the uncut control
   • 1 µl XbaI to X digest
   • 1 µl PstI to P digest
   • 1 µl XbaI and 1 µl PstI to X/P digest
6. Flick tube to mix and pulse spin
7. Incubate at 37°C for 15 minutes ("dry" heat block)

Agarose gel analysis of restriction digests

• Prepare a wide mini 1% agarose gel with a 20-well comb (use 150 ml) as on Day 1; two teams will share one gel
• Pulse spin the digest reactions and add 5 µl 6X LB to each
• Load ALL of each reaction on the gel (pipet slowly and carefully -- the wells will be full)
• Load 10 µl NEB Quick-Load 1 kb DNA Ladder
• Run the gel at 130 V for 30 minutes
• Photograph the gel and compare the observed bands to the standards
  
  Expected products:
  
  
  BBa_I13521 (strong constitutive production of red fluorescent protein (RFP), in pSB1A2, AmpR)
  • Uncut control = supercoiled (migrates farther than true size)
  • X digest = linearized/~3000bp
  • P digest = linearized/~3000bp
  • X/P digest = 2079 bp, 923 bp

Estimation of DNA amounts by absorbance at 260 nm

1. Turn on a spectrophotometer and let it warm-up for 5-10 minutes
2. Set the wavelength to 260 nm
3. Pipet 95 µl nuclease-free water into a disposable UV cuvette
4. "Zero" the instrument
5. Add 5 µl mini prep DNA to the 95 µl water; cover cuvette with parafilm and gently invert to mix
6. Read the absorbance
   Make sure that the cuvette is facing the SAME direction as for the blank
7. Calculate the DNA concentration for your sample (remember to account for dilution of the DNA)

Restriction digests of BioBrick™ plasmids

You will perform double digests on two BioBricks (BBa_R0040 and BBa_E0840); you will also set up an uncut control for each plasmid.   On day 3, you will gel purify the DNA and perform ligation and transformation.

NOTE: one set of digest reactions per team

1. Put 5 µl plasmid DNA (mini prep) in each tube (two tubes for each plasmid)
2. Add 13 µl nuclease-free water to each reaction (for a final volume of 25 µl)
3. Add 2.5 µl 10X NEB buffer #2 (BBa_R0040) OR #3 (BBa_E0840)
4. Add 2.5 µl 10X BSA to each tube
5. Add
   • 1 µl SpeI and 1 µl PstI to BBa_R0040
   • 1 µl XbaI and 1 µl PstI to BBa_E0840
   • 2 µl nuclease-free water to both uncut controls
6. Flick tube to mix and pulse spin
7. Incubate at 37°C for 15 minutes ("dry" heat block)
8. Pulse-spin all samples
9. Heat inactivate restriction enzymes at 80°C (oven) for 20 minutes
10. Store digests at -20°C (in a benchtop cooler) until day 3

Efficiency of Transformation (EOT) = # colonies / per µg of DNA
EOT is calculated by counting the number of colonies that grow on selective media following transformation and dividing by the total µg DNA used in the transformation. Dilutions must be calculated to determine the amount of DNA present in the volume of transformed culture placed on each plate: assume that the initial concentration of the plasmid DNA was 0.1 µg/µl; you added 1 µl of plasmid DNA to 40 µl electrocompetent cells.

Copyright, Acknowledgements, and Intended Use
Created by B. Beason (bbeason@rice.edu), Rice University, 21 November 2007
Updated 23 October 2009