20.109(F09): TA notes for module 2

Before module begins

Strains

• Streak out NB5 (= b-gal overproducing strain) on LB+Amp100
• Streak out NB188 (= strain for transformation/library screen) on LB+Amp100
• Streak out NB 334 (= bacterial photography strain) on LB+Cam34+Amp100
• prepare electroporation competent cells of NB188. Innoculate 3x 50 ml LB+Amp100 with 250 ul of an overnight culture of NB188 that was grown in LB+Amp100. For F09: grown in 125 ml flask in room temp shaker starting at 5PM and harvested at 10AM next day. Spin in clinical centrifuge 3.2K 5' and then spin sup down again. Resuspend cells from each 50 ml culture in cold 10% glycerol and move to 10 epps (1 ml each). Pellet. Wash cells with 1 ml cold 10% glycerol. Pellet. Wash cells with 250 ul cold 10% glycerol. Resuspend cells in 50 ul cold 10% glycerol and freeze -80° <6 months. Test electroporation before module starts.

Materials

• check that K+ and P+ library is available
• check stocks for protein gels/western including 12CA5 antibody, Epicentre prot ext'n soln, detection kit, kaleidoscope markers, gels, buffers

Day 1

• Test in Solid Media
• Test in Liquid Media
• Practice b-gal

Day before lab

• Set up 2.5 ml overnights of NB5 and NB334 (one culture/group)-->37° in LB+Amp100
• Make at least 500 ml of Z-buffer
  • for 500 ml
    • 8.05 g Na₂HPO₄*7H₂0
    • 2.75 g NaH₂PO₄*H₂O
    • 0.375 g KCl
    • 0.123 g MgSO₄*7H₂0
  • Dissolved in 500 ml H₂0 final volume
• Make 100 ml 4 mg/ml ONPG in Z-buffer, aliquot 1 ml/group and freeze rest
• Make 250 ml 1 M Na2CO3 in water, aliquot 5 ml/group and leave rest at RT
• Make 0.1% SDS (can dilute 10% solution), aliquot 0.5 ml/group and leave rest at RT

Day of lab

• Autoclave 50 ml of photography media/group. Not sure what the best way to do this will be. Perhaps want to autoclave large-ish batch of media and then aliquot to 100 ml bottles that have been autoclaved with a stirbar.
  • per 50 ml need:
    • 0.5g Tryptone
    • 0.25g Yeast Extract
    • 0.5g NaCl
    • 15mg Sgal
    • 25mg ferric ammonium citrate
    • 0.5g Low Melting Point Agarose
  • autoclave 30', cool to 42° in waterbath

Day 2

• Light/Dark b-gal
• Bacterial Photograph
• Oral Presentation Instruction in lab

Day of lab

• see day 1 info
• also need quiz

Day 3

• Transform Library
• Registry of Std Biological Parts
• Electronics

In advance

• Pour LB+Cam34+Amp100 petri dishes, need 2/group.
• Check stock of cuvettes, SOC media
• pre-run electroporation to check on best volume of cells for students to plate

Day of lab

• Each pair needs ice bucket, electroporation cuvette, 2 LB+Cam34+Amp100
• Front bench needs at least 2x electronics lab set up (see protocols)
• Gel running bench can have 2 electroporation machines set up
• In ice bucket on front bench: thaw K+ and P+ library

Day 4

• MUG to identify candidates
  • Restreak and set up ONs

In advance of lab

Make sure Top Agar is available, as well as sterile small tubes and a few aliquots of MUG (10 mg/ml in DMSO)

Day of lab

• Melt top agar and leave in 50° waterbath.
• Set up pipetaids and 5 ml pipets and small sterile tubes near waterbaths (front bench)
• Set out long wave UV lamps.

Day 5

• Protein Gel
• Miniprep/Digest/Agarose gel/Send to Sequence
• Set up Dark/Light ONs

Day 6

• Probe Western
• b-gal
• Seq analysis

Day 7

• "only" Journal Club