Team:University of Chicago/Notebook

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Home	Team	Project	Parts	Modeling	Notebook	Meetings	Papers

Lab Calendar

Click to see what lab work was done each day!

Individual Lab Notebook Pages

Damon Wang's notebook
Parijata "Jata" notebook
Nora Yucel's notebook
Laura Stone's notebook
Rob McConeghy's notebook
Daniel M. Choi's notebook

If you did it, record it

Resources

[http://www.freewebs.com/genehackers/ Team Website (External Link)]

Recipes

Media

LB Broth (1 L)

1 L dH20
10 g Bacto tryptone
5 g Bacto yeast extract
5 g NaCl
1 mL 2N NaOH
12 g agar (optional)
- 3 g agar + 250 mL broth per bottle
Autoclave

Note: Add 3 mL 1 M CaCl2 before phage transductions

PYE

This is a general-purpose Caulobacter medium and is typically used for plates and starter cultures from the freezer and for colonies off plates. Plates are incubated at 30C. Caulobacter colonies typically take 2-3 days to appear.

Per liter,

2g bacto-peptone
1g yeast extract
200mg magnesium sulfate heptahydrate
100mg calcium chloride dihydrate

Use deionized or distilled water; occasionally problems arise with less pure sources.

M13 Sweet

This is our production medium.

After autoclaving a liter of de-ionized water in a flask, add

40mL 25X M13 concentrate
10mL Hutner's Mineral Base Concentrate

We sometimes add a little extra calcium chloride (100mM stock to a final concentration of 1mM) to the flasks to encourage aggregates.

When the 1X M13 Sweet medium is assembled as indicated above, the final concentrations are:

0.5% glucose
0.15% glutamate
2mM phosphate
5 mM imidazole
0.04% ammonium chloride
1% Hutner's Mineral Salts

25X M13 concentrate

Component stock solutions:

125.0g glucose (or 137.5g glucose monohydrate) in a total volume of 600mL dH₂O
47.7g L-glutamic acid sodium salt (MW 187.13) in a total volume of 200mL dH₂O
8.5g imidazole in a total volume of 100mL dH₂O
3.4g KH₂PO₄ in a total volume of 50mL dH₂O
10g ammonium chloride in a total volume of 50mL dH₂O

All component stocks should be made in separate bottles, autoclaved, and once they are cooled, combined into a single sterile bottle. If the components are mixed first and then autoclaved, the stock turns black. Store the stock at room temperature or at 4C. Don't worry if after a few weeks the 25X stock turns color from clear-yellow to yellow-brown.

SOB Medium

Used in growing bacteria for preparing chemically competent cells

Ingredients

0.5% (w/v) yeast extract
2% (w/v) tryptone
10 mM NaCl
2.5 mM KCl
20 mM MgSO4

Per liter:

5 g yeast extract
20 g tryptone
0.584 g NaCl
0.186 g KCl
2.4 g MgSO4

Note: Some formulations of SOB use 10 mM MgCl2 and 10 mM MgSO4 instead of 20 mM MgSO4.

SOB medium is also available dry premixed from Difco, 0443-17.
Adjust to pH 7.5 prior to use. This requires approximately 25 ml of 1M NaOH per liter.

==== SOC MediuProxy-Connection: keep-alive Cache-Control: max-age=0

(per 100 ml) ==== Note This medium should be prepared immediately before use.

Add 2 ml of filter-sterilized 20% (w/v) glucose or 1 ml of filter-sterilized 2 M glucose to 100 ml sterile SOB

Buffers

CCMB80

10 mM KOAc pH 7.0 (10 ml of a 1M stock/L)
80 mM CaCl2.2H2O (11.8 g/L)
20 mM MnCl2.4H2O (4.0 g/L)
10 mM MgCl2.6H2O (2.0 g/L)
10% glycerol (100 ml/L)
adjust pH DOWN to 6.4 with 0.1N HCl if necessary
- adjusting pH up will precipitate manganese dioxide from Mn containing solutions.
sterile filter and store at 4°C
slight dark precipitate appears not to affect its function

Note: Create 1M concentration stocks to make buffer prep easier in future.

CCMB80: with aqueous stocks

10 mM KOAc pH 7.0: 10mL of 1M stock/L
80 mM CaCl2.2H2O: 80mL/L
20 mM MnCl2.4H2O: 20mL/L
10 mM MgCl2.6H2O: 10mL/L
10% glycerol: 100 ml/L

Hutner's Mineral Base Concentrate

It is not difficult, but follow directions carefully.

20g nitrilotriacetic acid (NTA)
54.5g magnesium sulfate heptahydrate
6.67g calcium chloride dihydrate
18.5mg ammonium molybdate tetrahydrate
198mg iron sulfate monohydrate (or 323.7mg of the heptahydrate)
100mL Metals-44

TO BE CONTINUED

5x Ligation Adjustment Buffer

Intended to be mixed with ligation reactions to adjust buffer composition to be near the CCMB80 buffer
KOAc 40 mM (40 ml/liter of 1 M KOAc solution, pH 7.0)
CaCl2 400 mM (200 ml/l of a 2 M solution)
MnCl2 100 mM (100 ml/l of a 1 M solution)
Glycerol 46.8% (468 ml/liter)
pH adjustment with 2.3% of a 10% acetic acid solution (12.8ml/liter)
- Previous protocol indicated amount of acetic acid added should be 23 ml/liter but that amount was found to be 2X too much per tests on 1.23.07 --Meaganl 15:50, 25 January 2007 (EST)
water to 1 liter
autoclave or sterile filter
Test pH adjustment by mixing 4 parts ligation buffer + 1 part 5x ligation adjustment buffer and checking pH to be 6.3 - 6.5
Reshma 10:49, 11 February 2008 (CST): Use of the ligation adjustment buffer is optional.

DNA and protein gels

Agar

0.8%

Weigh out .8g agarose
Put agarose in 500mL Erlenmeyer flask.
Add 100mL 1X TBE Buffer. Swirl.
Microwave at 100% power until the agarose starts to dissolve. Every 30-40 seconds, stop and swirl. Continue microwaving until solution boils and agarose is dissolved.
After agarose is dissolved, cover flask with foil or aran wrap and place in 55-60C waterbath until needed.
When Agarose has cooled to enough to touch comfortably, add 10microliters Syber Safe. Swirl.
Pour 30mL gels. Use 50Ml conical tube to measure 30mL.

1.2%

Same as above but use 1.2 g agarose instead of .8g

Coomassie Blue Stain/Destain Recipe

450 ml water
450 ml methanol
100 ml 100% (glacial) acetic acid

Add 2 g per liter coomassie blue for stain. Destain is exactly the same, except do not add coomassie blue.
To stain the gel, incubate it with gentle shaking for at least 30 minutes. Pour the stain back into the bottle (you can reuse it). Rinse out the residual stain with a small amount of destain solution, and then incubate with gentle shaking for ~1 hour or more. To speed up destaining, you can put some folded KimWipes in the dish. The dye will adsorb to these, which will remove it from solution.
After your gel is destained, you can rehydrate it in distilled water and then image it and/or place it on a piece of filter paper and dry it with a gel dryer.

Protocols

Growing Cells

TOP10 Competent Cells

Prechill plasticware and glassware

Preparing seed stocks

Streak TOP10 cells on an SOB plate and grow for single colonies at 23°C
- room temperature works well
Pick single colonies into 2 ml of SOB medium and shake overnight at 23°C
- room temperature works well
Add glycerol to 15%
Aliquot 1 ml samples to Nunc cryotubes
Place tubes into a zip lock bag, immerse bag into a dry ice/ethanol bath for 5 minutes
- This step may not be necessary
Place in -80°C freezer indefinitely.

Preparing competent cells

Inoculate 250 ml of SOB medium with 1 ml vial of seed stock and grow at 20°C to an OD600nm of 0.3
- This takes approximately 16 hours.
- Controlling the temperature makes this a more reproducible process, but is not essential.
- Room temperature will work. You can adjust this temperature somewhat to fit your schedule
- Aim for lower, not higher OD if you can't hit this mark
Centrifuge at 3000g at 4°C for 10 minutes in a flat bottom centrifuge bottle.
- Flat bottom centrifuge tubes make the fragile cells much easier to resuspend
- It is often easier to resuspend pellets by mixing before adding large amounts of buffer
Gently resuspend in 80 ml of ice cold CCMB80 buffer
- sometimes this is less than completely gentle. It still works.
Incubate on ice 20 minutes
Centrifuge again at 4°C and resuspend in 10 ml of ice cold CCMB80 buffer.
Test OD of a mixture of 200 _l SOC and 50 _l of the resuspended cells.
Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test.
Incubate on ice for 20 minutes
Aliquot to chilled screw top 2 ml vials or 50 _l into chilled microtiter plates
Store at -80°C indefinitely.
- Flash freezing does not appear to be necessary
Test competence (see below)
Thawing and refreezing partially used cell aliquots dramatically reduces transformation efficiency by about 3x the first time, and about 6x total after several freeze/thaw cycles.

Measurement of competence

Transform 50 _l of cells with 1 _l of standard pUC19 plasmid (Invitrogen)
- This is at 10 pg/_l or 10-5 _g/_l
- This can be made by diluting 1 _l of NEB pUC19 plasmid (1 _g/_l, NEB part number N3401S) into 100 ml of TE
Hold on ice 0.5 hours
Heat shock 60 sec at 42C
Add 250 _l SOC
Incubate at 37 C for 1 hour in 2 ml centrifuge tubes rotated
- using 2ml centrifuge tubes for transformation and regrowth works well because the small volumes flow well when rotated, increasing aeration.
- For our plasmids (pSB1AC3, pSB1AT3) which are chloramphenicol and tetracycline resistant, we find growing for 2 hours yields many more colonies
- Ampicillin and kanamycin appear to do fine with 1 hour growth
Plate 20 _l on AMP plates using sterile 3.5 mm glass beads
- Good cells should yield around 100 - 400 colonies
- Transformation efficiency is (dilution factor=15) x colony count x 105/µgDNA
- We expect that the transformation efficiency should be between 5x108 and 5x109 cfu/µgDNA

Glycerol Stocks

Materials

40% glycerol solution
Cryogenic vials

Method

Add 1 ml of 40% glycerol in H2O to a cryogenic vial.
Add 1 ml sample from the culture of bacteria to be stored.
Gently vortex the cryogenic vial to ensure the culture and glycerol is well-mixed.
- Alternatively, pipet to mix.
Use a tough spot to put the name of the strain or some useful identifier on the top of the vial.
On the side of the vial list all relevant information - part, vector, strain, date, researcher, etc.
Store in a freezer box in a -80C freezer. Remember to record where the vial is stored for fast retrieval later.

Notes

While it is possible to make a long term stock from cells in stationary phase, ideally your culture should be in logarithmic growth phase. Prof. Schonbaum's version

Grow a 3 ml culture overnight
Add 200 µl 100% glycerol to 1 ml cells in a cryotube (final concentration should be 15-20% glycerol different concentrations from different protocols
Mix by vortexing and let sit for a little bit. Then put the tubes in the -80°C freezer.

@@ Line 162: / Line 162: @@
 ===== 1.2% =====
 Same as above but use 1.2 g agarose instead of .8g
+====Coomassie Blue Stain/Destain Recipe====
+ml water<br>
+ml methanol<br>
+ml 100% (glacial) acetic acid<br><br>
+#Add 2 g per liter coomassie blue for stain.  Destain is exactly the same, except do not add coomassie blue.
+#To stain the gel, incubate it with gentle shaking for at least 30 minutes.  Pour the stain back into the bottle (you can reuse it).  Rinse out the residual stain with a small amount of destain solution, and then incubate with gentle shaking for ~1 hour or more.  To speed up destaining, you can put some folded KimWipes in the dish.  The dye will adsorb to these, which will remove it from solution.
+#After your gel is destained, you can rehydrate it in distilled water and then image it and/or place it on a piece of filter paper and dry it with a gel dryer.
 == Protocols ==