20.109(S11):Complete DNA design (Day2)

Introduction

Protocols

Part 1: Complete DNA design

Part 2: Test liquid cultures for B-gal production

FROM NK, IN REVISION BUT MOSTLY UPDATED

With this assay you will determine the amount of beta-galactosidase activity associated with your cultures from last time. A table is included here to help you organize your assay, but you can make one of your own if you prefer. You may want to make an Excel sheet and print it out instead...

The sample order below is recommended in order to minimize the risk of saturating samples expected to produce a lot of B-gal. [Or have them figure out/explain in hw?]

When you first try this assay you may find 15 second intervals too fast, and when you are expert at it you may find them too slow. Somewhere between 10 and 20 seconds should be a good time interval to shoot for.

Tube #	Sample	OD600	Time started	Time stopped	Time elapsed (calculated)	OD420	OD550	Units (calculated)
0	blank		0:00
1	IPTG-1		0:15
2	IPTG-2		0:30
3	IPTG/AHL-1		0:45
4	IPTG/AHL-2		1:00
5	neg-1		1:15
6	neg-2		1:30
7	AHL-1		1:45
8	AHL-2		2:00

1. Retrieve the cultures that you prepared last time, and prepare X mL of a 1:10 dilution of each one.
   • Use 0.6 mL of the dilution to measure the OD₆₀₀ for each sample.
   • Save the rest for the B-gal assay.
2. Add 400 μl of Zbuffer to 9 eppendorf tubes labeled 0-8.
3. Add 100 μl of the diluted cells to each tube. See chart above for guidance. Add 100 ul of Zbuffer to tube 0, to serve as your blank.
   • POSSIBLY EVEN WANT TO GO LOWER - 50 μL and 450 of Z-buffer. [For 5 hr assays, 1:5 dilution even would be fine or 20 uL/480 uL directly]
4. Next you will lyse the cells by add 20 μl of 0.1% SDS to each eppendorf.
5. To more fully lyse the cells, you should also add 30 μl of chloroform (CHCl₃) to each tube. Do this in the hood since chloroform is volatile and toxic.
   • You will need to hold the pipet tip close to the eppendorf as you move between the chloroform stock bottle and your eppendorfs since chloroform has a low surface tension and will drip from your pipetmen. Be sure to dispose of your pipet tips in the chloroform waste container located on the right side of the hood.
6. To complete the cell lysis, vortex the tubes for 10 seconds each. You should time these precisely since you want the replicates to be treated as identically as possible.
   • You should be able to fit 2-3 tubes on the vortex at once.
7. Start the reactions by adding 100 μL of ONPG to each tube at 10 second intervals, including your blank. Invert to mix.
8. Stop the reactions by adding 250 μL of Na₂CO₃ to each tube once sufficient yellow color has developed.
   • “Sufficient” is defined as yellow enough to give a reliable reading in the spectrophotometer, best between 0.3 and 1.0. It is about the amount of yellow color that you see for the yellow tips for your P200. Keep in mind, though, that adding Na₂CO₃ makes the reactions more yellow!
   • Try to stop the duplicate reactions at equal intervals and be sure to note the time you are stopping the reactions.
9. When all your samples have been stopped, add 250 μL of Na₂CO₃ to the blank and spin all the tubes in the microfuge for 1 minute at 13,000 RPM to pellet any cell debris.

Option 1

1. Move 0.7 ml of each reaction to plastic cuvettes.
   • Avoid the chloroform that will be at the bottom of your tubes. If you add the chloroform to the cuvettes, it will "etch" the cuvette windows and mess up your readings.
2. Read the absorbance at 420nm. These values reflect the amount of yellow color in each tube.
3. Read the absorbance of each at 550 nm. These values reflect the amount of cell debris and differences in the plastic cuvettes themselves.

Option 2

1. Move 0.2 ml of each reaction to a 96-well plate.
   • Avoid the chloroform that will be at the bottom of your tubes.
2. Go to BPEC with a member of the teaching faculty to automatically read your samples at 420 and 550 nm.

1. Dispose of your samples properly: liquid contents of cuvettes/microplates can go down the sink, empty cuvettes can go in the sharps containers, and CHCl₃ can go into a waste bottle in the hood.
2. The β-gal activity in each sample is reported as "Miller Units" according to the following formula:

1 Miller Unit = $ 1000 * \frac{(Abs{420} - (1.75*Abs{550}))}{(t * v * OD{600})} $

where:
Abs 420 is a measure of the yellow color produced by the β-gal activity.
Abs 550 is a measure of cell debris.
OD 600 is a measure of the cell density.
t is the reaction time from start to stop, measured in minutes.
v is the culture volume that you added to the reactions, measured in mls.
You can average the duplicate values for each sample unless you know of a reason not to (e.g. one tube spilled or had the incorrect amount of something added to it...)

Part 3: Observe solid cultures

For next time

==Reagent list==