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

In advance of lab

• Streak out strains NB5 (= b-gal overproducing strain) on LB+Amp100
• Streak out NB334 (= bacterial photography strain) on LB+Cam34+Amp100
• Autoclave large and small tubes if needed

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

• One box of cuvettes/team
• 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 except don't need NB5 overnight cultures
• 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, EP cells just before use
• Front bench also needs alcohol burners, EtOH beakers, spreaders, strikers

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

• NO quiz
• Melt top agar and leave in 50° waterbath.
• Set up pipetaids and 5 ml pipets and vortexers 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

In advance

• check levels on solutions for protein gel (TGS, TBS, Tween, milk powder)
• make Transfer Buffer and store in delicase (4°)
  • 3.03 g Trizma base
  • 14.4g glycine
  • 200 ml methanol
  • to 1L with good H₂O
  • Store at 4°C
• aliquot miniprep solutions (see below)

Day of lab

• Need quiz
• Make 800 ul of Epicentre "EasyLyse" solution for each group. This is done by mixing (in the following order):
  • 0.4 ml sterile H₂O
  • 1.6 ul 1M MgCls (kept in Epicentre kit at RT)
  • 0.4 ml Lysis Buffer (kept in Epicentre kit at RT)
  • 0.8 ul "enzyme mix" (kept in tote in -20°)
  • mix just before lab and leave on ice until students request it
• Make TBS+T+milk (25 ml/group)
  • TBS-T: Dilute 100 ml 10X TBS with 900 ml H20 then add 10 ml 10% Tween20
  • TBS-T + 5% milk: add 2.5g milk powder to 50 ml TBS-T. Mix on stir plate or in conical at 37° on nutator until milk dissolved
• Pour 1 agarose gel, 1% in TAE (100ml) + 2 ul EtBr each. Use 2 10-tooth combs/gel.
• Each group needs miniprep solutions (Solutions 1, 3, NaOH and SDS) for each group so stocks don't get contaminated.
  • 400 ul Solution I
  • 500 ul SDS 2%
  • 500 ul 0.4M NaOH
  • 600 ul Solution III
  • 4 ml 100% Ethanol
  • 2 ml 70% Ethanol
  • sterile water bottle for each group
• Near end of lab, assemble cocktail of NdeI and MluI in NEB3. For 20X cocktail, mix 250 ul H₂O, 50 ul 10X NEB3, 5 ul each enzyme. Add 15 ul of cocktail to 10 ul of student's samples, incubate 30 minutes at 37° and run on agarose gel with 1KB marker lane. Photograph and post.
• Near end of lab, thaw sequencing primer NO289

Day 6

• Probe Western
• b-gal
• Seq analysis

Day of lab

• Need quiz
• see b-gal reagents, day 1

Day 7

• No quiz
• "only" Journal Club

Recipes/Reagents

Growth media

1. LB: 10 g Tryptone, 5 g Yeast Extract, 10 g NaCl per liter. 20g of Agar for plates. Autoclave 30 minutes with stirbar. Pour when ~55°. Let plates dry ON on bench and store in sleeves in 4°. For LB+ antibiotics plates, add the antibiotics after autoclaving, once the mixture has cooled down.
2. Top Agar: 10 g Tryptone, 5 g Yeast Extract, 10 g NaCl, 1 g MgCl2*6H20 7 g Agar per liter. Autoclave then aliquot to 50 ml conical tubes or sterile bottles. Store at RT. Melt in microwave in beaker of water, 2’ then keep molten in 55° water bath.
3. Amp: 100 mg/ml in H20. Filter and store at 4°. Use at 1:1000
4. Cam: 34 mg/ml in EtOH. No need to filter sterilize. Use at 1:1000

DNA Miniprep

1. Soln I for miniprep: 2.3 ml 40% glucose, 2.5 ml 1M Tris 8, 2 ml 0.5M EDTA. To 100 ml with good H20. Store at RT
2. Soln II for miniprep: equal parts 2% SDS (2g/100 ml H20): 0.4M NaOH (1.6g/100 ml H20). Store components at RT. Mix just enough just before using.
3. Soln III for miniprep: 29.4 g KAc dissolved in 60 ml H20. Add 11.5 ml glacial acetic acid. Bring to 100 ml final volume. Store at RT.

Agarose Gel

1. DNA gel: 1% agarose gel in 1X TAE, 1 g agarose, 100mL 1X TAE, 2 ul EtBr (wear nitrile gloves when handling EtBr!)
2. Loading dye for agarose gel: 250 ul 1% XC (xylene cyanol), 750 ul 40% glycerol, 10 ul RNase. Store at RT.
3. 1kb marker: 10uL 1kb marker stock (in -20 freezer), 10uL loading dye, 90uL H20

Western Blot

1. 2X sample dye for protein gel (no BME): 4 ml 10% SDS, 5 ml 40% glycerol, 1 ml 1M Tris 6.8, 0.5 ml <1% bromophenol blue, stocks on NK's bench
2. 1X sample dye for protein gel using Sigma mix: 500 ul 2X sample dye, 200 ul H2O, 200 ul 10% SDS, 100 ul BME
3. Transfer buffer: 3.03 g Trizma base, 14.4g glycine, 200 ml methanol, to 1L with good H2O. Store at 4°C
4. TBS-T: Dilute 100 ml 10X TBS with 900 ml H20 then add 10 ml 10% Tween20
5. TBS-T + 5% milk: add 2.5g milk powder to 50 ml TBS-T. Mix on stir plate or in conical at 37° on nutator until milk dissolved