20.109(F11): TA notes for module 2

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Before module begins

Current module
Archive of TA notes
Fall 2009
Fall 2010,

Notebook grading checklist


  • Streak out NB5 (= b-gal overproducing strain) on LB+Amp100
  • Streak out NB462 (= strain for transformation/library screen) on LB+Amp25 (can also spread plate with 25 ul Kan10 to check strain)
  • Streak out NB466 (= bacterial photography strain, alternative = NB334) on LB+Cam34+Amp25+Kan10
  • Streak out NB467 and NB468 (= kinase dead strain H557A, in XL1-blue and photography strain respectively). NB467 should be streaked out on LB+Cam34 and NB468 on LB+Amp25+Cam34+Kan10. If there is some problem with this "kinase dead" pair of strains, then it's also possible to use I566V mutant, NB469 and NB470.
  • prepare electroporation competent cells of NB462. Innoculate 3x 50 ml LB+Amp25+Kan10 with 50 ul of an overnight culture of NB462 that was grown in LB+Amp25+Kan10. For F11: grown in 125 ml flask in room temp shaker starting at 3PM and harvested at 9AM next day. Pool cultures in one flask to make uniform (measured OD600 = 0.515). Divide between 4 50 ml conical tubes of ~35 ml each to spin in clinical centrifuge 3.2K 5' and then spin sup down again. Resuspend cells from each 50 ml conical in 10 ml cold 10% glycerol (40 ml total) and move 20 ml to 20 epps (1 ml each). Pellet. Add other 20 ml to those epps. Pellet again. 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 for a total of 20 epps of EP comp tubes and freeze -80° <6 months. Test electroporation before module starts.
  • miniprep H557A kinase dead control plasmid from NB467 and/or I566V mutant from NB469. These will be used for electroporation/screening controls on Day 3.


  • check that K-P+ library is available
  • check stocks for protein gels/western including anti-H6EnvZ antibody, Epicentre prot ext'n soln, detection kit, kaleidoscope markers, gels, buffers

Day 1

Lab has 3 parts:

  • 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 NB466 (= bacterial photography strain) on LB+Cam34+Amp25+Kan10
  • Autoclave large and small tubes if needed

Day before lab

  • Set up 2.5 ml overnights of NB5 (one culture/group)-->37° in LB+Amp100
  • Set up 2.5 ml overnights of NB466 (one culture/group)--> 37° in LB+Cam34+Amp25+Kan10
  • 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
  • Sleeve of empty petri dishes (at least 2 needed per team)
  • Autoclave 30 ml of photography media/group. Best to make media in batches of 100 ml using 250 ml bottles with stir bars. Each bottle will provide enough for three groups. Pre-heat 42° waterbath!
    • per 100 ml need:
      • 0.5g Tryptone
      • 0.25g Yeast Extract
      • 0.5g NaCl (can use 100 ml of pre-made LB instead of these first 3 ingredients if there is lots of LB in the lab)
      • 30mg Sgal
      • 50mg ferric ammonium citrate
      • 1 g Low Melting Point Agarose (be sure to use low melt point agar!)
    • autoclave 30', stir at RT 3-5', cool to 42° in waterbath on instructor's bench at front of lab

Day 2

Lab has 4 parts:

  • Light/Dark b-gal
  • Bacterial Photograph
  • TinkerCell Model and Simulation
  • Oral Presentation Instruction in lab

Day of lab

  • see day 1 info except
    • don't need NB5 overnight cultures. Students will use light/dark grown cultures they've grown.
    • need twice as much of each b-gal reagent since students are likely to run 12 not 6 reactions.
    • each group will only run one bacterial photography plate, not two, so can divide volume in half.
  • also need quiz
  • each bench will need a copy of the print-out of the tinkercell tutorial
  • front bench needs transparencies for printing.

Day 3

Lab had 4 (maybe 4) parts:

  • Electroporate Library
  • Registry of Std Biological Parts
  • Electronics
  • TinkerCell Simulations, if the students were not done on Day 2

In advance

  • Pour Tetrazolium+Cam34+Amp25+Kan10 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
  • Miniprep kinase dead mutants to electroporate as controls

Day of lab

  • Each pair needs ice bucket, electroporation cuvette, 2 Tetrazolium+Cam34+Amp25+Kan10
  • 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-P+ library, EP cells just before use
  • Front bench also needs alcohol burners, EtOH beakers, spreaders, strikers

Day 4

  • Students will identify 2 candidates from K-P+ library.
    • TA needs to restreak for single colonies on the first day between labs, then
    • set up 2.5 ml overnights in light and dark from single colony on the second night between labs.

This is a lot of work!

Day of lab

  • NO quiz
  • When students identify two candidate, you can restreak them onto LB+Cam34+Amp25+Kan10 to grow 37° overnight.
  • One day before the next lab you can set up overnight cultures in LB+Cam34+Amp25+Kan10 in the light and dark. Will also want to set up unmutated and kinase dead controls (NB466 and NB468 and/or NB470)

Day 5

Lab has 3 parts:

  • DNA: miniprep/send to sequencing facility
  • Protein Activity: b-gal from light and dark
  • Writing instruction

In advance

  • ONE DAY BEFORE LAB: will need to set up overnight cultures of NB466 (bacterial photography strain), NB468 (kinase dead bacterial photography strain) and the 2 mutants that the students have selected. A 5 ml overnight in LB+Cam34+Amp25+Kan10 for each should be enough. Split the 5 ml between two snap-cap tubes and grow them in the light/dark, overnight. May want to do the controls in advance to manage the work better.
  • aliquot miniprep solutions (see below)

Day of lab

  • Need quiz
  • 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, thaw sequencing primer NO296 and dilute 1:20 in water. Each group needs about 10 ul of dilute oligo. Also need a few 8 strip PCR tubes to send to seq.
  • Each group also needs b-gal assay solutions, like Day 1 of the module.
    • Z-buffer, 10 ml/ group
    • 4 mg/ml ONPG in Z-buffer, aliquot of 1.5 ml/group
    • 1 M Na2CO3 in water, aliquot of 7.5 ml/group
    • 0.1% SDS in water, aliquot of 0.5 ml/group
    • Box of cuvettes
  • Each group will need to set up 5 ml overnight cultures from their dark grown mutants as well as wild type (total of 3 tubes/team). These overnights will be grown in sterile tubes. You can put out a rack of tubes as well as a 100ml bottle of LB with LB+Cam34+Amp25+Kan10 already added. You should also put out a sleeve of 5 ml pipets. They will need to dispense then media to the tubes and then add 5 ul of their overnights to start these new cultures.

End of lab

  • Store remainder of overnight cultures for Day 6 in case students want them
  • Put student overnight cultures on the culture wheel at 37°
  • Drop samples in Genewiz box

Day 6

  • Photograph
  • Seq analysis
  • Protein Gel/blot
  • Possible additional experiments as dictated by the students

In advance of lab

  • check levels on solutions for protein gel (TGS, TBS, Tween, milk powder)
  • make Transfer Buffer and store in delicase (4°)
    • NEED 1 liter/pair of blots (= 4 teams).
    • 3.03 g Trizma base
    • 14.4g glycine
    • 200 ml methanol
    • to 1L with good H2O
    • Store at 4°C must be cold on day of lab

Day of lab

  • Need quiz
  • Prepare 2X SB
    • 500 ul Sigma dye G2526
    • 200 ul H2O
    • 200 ul 10% SDS
    • 100 ul BME
  • Dilute purified H6-EnvZ protein and mix with 2xSB (2.5 ul into 100 ul 2xSB). Students will need 40 ul/team so aliquot 50ul for each.
  • 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
    • Transfer buffer (1 liter /tank). One tank holds two gels, i.e. 4 groups.
  • Set up acrylamide gel in 1X TGS just before lab. If there are extra old gels that can be used to practice loading, set one up on the white team bench or on the end of the gel running bench.

Day 7

  • Probe Western
  • Need quiz

In lab

  • Bring blots from fridge to RT just before lab starts
  • Just before lab thaw primary antibody to His6-EnvZ. You will need 10 ul per group.
  • Make sure shaker platform in chemical hood is set to rotate at ~60 rpm.
  • Confirm that there is sufficient GAR-AP antibody (4° deli case, from Sigma)
  • Student groups will need ~250 ml TBS-T each.
  • Aliquot 1X development solution so 25 ml conical available for each group.
  • Bring out development kit just as needed.
  • Once gels developed, scan or photograph and post images to talk page of today's lab.

Day 8

  • No quiz
  • "only" Journal Club


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. Tetrazolium indicator media: for 400 ml: 10.2 grams Antibiotic Medium #2, 20 mg Tetrazolium (kept in 4° delicase with chemicals). Add 380 ml H2O then heat at setting "5" in hood on stirplate to help dissolve agar. Autoclave 30 minutes and cool to ~55° then add 20 ml of 20% lactose (4g/20 ml H20, need to heat this to dissolve then filter sterilize) and 400 ul Amp25 + 400 ul Cam34 + 400 ul Kan10. Let plates dry ON on bench and store in sleeves in 4°. Plates may be used for ~1 month.
  3. Amp: 25 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
  5. Kan: 10 mg/ml in H20. Filter and store at 4°. 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