Interlab

D1: CalC2 Competent Cell Preparation

Introduction

Protocols adapted from: Here

Materials

• DH5α cells (2013 Genetics lab stock)
• 1 L liquid LB
• 2 LB plates
• Inoculating loop (yellow one)
• MgCl2
• CaCl2
• Glycerol
• 1.5 mL Eppendoft tubes
• Large rack + eppendorf rack
• Liquid nitrogen
• dH2O (deionized water)
• 3 Filter sterilization bottles
• Spectrophotometer
• Centrifuge for large tubes, at 4˚C (chem lab)
• Ice + bucket

Procedure

Preparing Solutions

MgCl2 solution (0.1 M)

1. 10.16 g MgCl2 in 500 ml dH20
2. Filter sterilize into bottle and label

CaCl2 solution (0.05 M)

3. 3.67 g CaCl2 in 500 ml dH20
4. Filter sterilize into bottle and label

CaCl2 w/ Glycerol solution (0.1 M CaCl2)

5. 2.94 g CaCl2 + 28 ml Glycerol + 172 ml dH20
6. Filter sterilize into bottle and label

Preparing cells

First day

7. Pick frozen DH5α cells and streak out on 2 LB Plates, place in 37˚C incubator overnight

Second day

8. Make sure there are good colonoes on the plates, tape and label, place in 4˚C fridge for use later

Third day

9. Pick a single colony from the plate and grow overnight in 5-10 ml of LB

Fourth day

10. Dilute the preparation 1:100 in LB (NO AMP): 5ml overnight culture into 500 ml LB
11. Grow the cells to Optical Density 600 = 0.6

After one hour, take OD reading, then take readings every 15 min, should take approx. 3 to 4 hrs.

12. Split the preparation into 2 large sterile centrifuge bottles
13. Centrifuge at 4K at 4˚C for 10 minutes
14. ON ICE: Pour off supernatant and redissolve each pellet in 125 ml 0.1 M MgCl2 (4˚C fridge)
15. ON ICE: Incubate the cells on ice for 7-10 minutes
16. Centrifuge again at 4K for 10 minutes
17. Dissolve each pellet in 125 ml of .05 M CaCl2 (Done in Cold Room)
18. Incubate on ice for 20 minutes (In Cold Room)
19. Centrifuge at 4K for 10 minutes @ 4˚C
20. Dissolve in 50 ml 0.1 M CaCl2 w/glycerol (In cold room)
21. Once dissolved, dispense 500 µL aliquots into 1.5ml eppendorf tubes, THAT HAVE BEEN PRE-CHILLED ON ICE.
22. Put on dry ice for 1.5 min before placing them in the -80˚C freezer

D2: Competent Cell Test

Introduction

The kit includes three vials of purified plasmid DNA from BBa_J04450 (RFP construct) in plasmid backbone pSB1C3. Each vial contains 50 µL of DNA at a different concentration: 100 pg/µL, 50 pg/µL, 10 pg/µL. Perform transformations with each of these to determine how efficient your competent cells are.

Protocols adapted from: Here

Materials

• DH5α cells (2013 Genetics lab stock)
• 70% ethanol
• Paper towels
• Lab marker / Sharpie
• 1.5 mL microcentrifuge tubes
• Container for ice + ice
• Competent cell aliquot(s)
• Competent Cell Test Kit
• Agar plates with chloramphenicol
• 42°C Waterbath (or hot water source and thermometer)
• 37°C Incubators (oven and shaker)
• SOC media
• Sterile glass beads or sterile cell spreader
• Pipettor + tips

Procedure

Estimated time: 30 minutes active, 1.5 hours incubation

1. Clean your working area by wiping down with 70% ethanol.
2. Thaw competent cells on ice. Label one 1.5 mL microcentrifuge tubes for each transformation and then pre-chill by placing the tubes on ice.

Do triplicates (3 each) of each concentration if possible, so you can calculate an average colony yield.

3. Spin down the DNA tubes from the Competent Cell Test Kit/Transformation Efficiency Kit to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient. Note: There should be 50 µL of DNA in each tube sent in the Kit.
4. Pipet 1 µL of DNA into each micro centrifuge tube.
5. Pipet 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
6. Incubate on ice for 30 minutes. Pre-heat waterbath now to 42°C. Otherwise, hot water and an accurate thermometer works, too!
7. Heat-shock the cells by placing into the waterbath for 45 seconds (no longer than 1 min). Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes.
9. Add 950 µL of SOC media per tube, and incubate at 37°C for 1 hour shaking at 200-300rpm. (Added floating boats and taped to the rack in the shaker), Prepare the agar plates during this time: label them and prepare spreader
10. Spin down the tubes at 10,000 rpm for 30 sec
11. Discard supernatant
12. Resuspend in 100 µL SOC and vortex to mix
13. Pipet 50 µL from each tube onto the appropriate plate, and spread the mixture evenly across the plate. Incubate at 37°C overnight or approximately 16 hours. Position the plates with the agar side at the top, and the lid at the bottom.

Next Day

14. Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation.

Efficiency (in cfu/µg) = [colonies on plate (cfu) / Amount of DNA plated (ng)] x 1000 (ng/µg)

*Note: The measurement "Amount of DNA plated" refers to how much DNA was plated onto each agar plate, not the total amount of DNA used per transformation. You can calculate this number using the following equation:

Amount of DNA plated (ng) = Volume DNA added (1 µL) x concentration of DNA (refer to vial, convert to ng/µL) x [volume plated (100 µL) / total reaction volume (1000 µL)]

Results:

15. Competent cells should have an efficiency of 1.5x10^8 to 6x10^8 cfu/µg DNA, where "cfu" means "colony-forming unit" and is a measurement of cells.
16. Here are some sample results:

DNA concentration 10 pg/µL 50 pg/µL 100 pg/µL

# of colonies 280 - 360 500 - 1000+ 1000+

D3: SOC Media Preparation

Introduction

Protocol adapted from Genetics Lab recipe book (Bernadette)

Materials

• 2% Tryptone
• 0.5 % Yeast Extract
• 10 mM NaCl
• 2.5 mM KCl
• 10 mM MgSO4 . 7H2O
• 10 mM MgCl2 . 6H2O
• 20 mM Glucose
• dH2O
• Autoclave
• 1 L measuring cylinder
• 1 L bottle

Procedure

1. Add the following:

	50 mL	100 mL
2% Tryptone	1 g	2 g
0.5% Yeast Extract	0.25 g	0.5 g
10mM NaCl	0.2922 g	0.5844 g
2.5 mM KCl	0.00932 g	0.0186 g
10 mM MgSO4.7H2O	0.123 g	0.246 g
10 mM MgSO4.6H2O	0.102 g	0.204 g
20 mM Glucose	0.18 g	0.36 g
dH2O	Up to volume	Up to volume

2. Autoclave

D4: Overall Protocol

Introduction

Protocol adpated from: Here

Materials

• Competent cells ( Escherichia coli strain DH5α)
• LB (Luria Bertani) media
• Chloramphenicol (stock concentration 25 mg/mL dissolved in EtOH - working stock 25 ug/mL) 50 ml Falcon tube (or equivalent, preferably amber or covered in foil to block light)
• Incubator at 37°C
• .5 ml eppendorf tubes for sample storage
• Ice bucket with ice
• Pipettes
• Devices (from InterLab Measurement Kit 7):
1. Positive control - well 21B
2. Negative control - well 21D
3. Test Device 1: J23101+I13504 - well 21F
4. Test Device 2: J23106+I13504 - well 21H
5. Test Device 3: J23117+I13504 - well 21J
6. Test Device 4: J23101.BCD2.E0040.B0015 - well 21L
7. Test Device 5: J23106.BCD2.E0040.B0015 - well 21N
8. Test Device 6: J23117.BCD2.E0040.B0015 - well 21P

Procedure

Workflow

Day 1 - Sunday - (23/7/2017) :

Transform Escherichia coli DH5α with these following plasmids (Distribution Kit plate 7):

Positive control (+ve control) - well 21B

Negative control (-ve control) - well 21D

Test Device 1: J23101+I13504 (TD1) - well 21F

Test Device 2: J23106+I13504 (TD2) - well 21H

Test Device 3: J23117+I13504 (TD3) - well 21J

Test Device 4: J23101.BCD2.E0040.B0015 (TD4) - well 21L

Test Device 5: J23106.BCD2.E0040.B0015 (TD5) - well 21N

Test Device 6: J23117.BCD2.E0040.B0015 (TD6) - well 21P

TRANFORMATION PROTOCOL USED:

1. Resuspend DNA in selected wells in the Distribution Kit with 10µl dH20. Pipet up and down several times, let sit for a few minutes. Resuspension will be red from cresol red dye.
2. Label 1.5ml tubes with part name or well location. Fill lab ice bucket with ice, and pre-chill 1.5ml tubes (one tube for each transformation, including your control) in a floating foam tube rack.
3. Thaw competent cells on ice: This may take 10-15min for a 260µl stock. Dispose of unused competent cells. Do not refreeze unused thawed cells, as it will drastically reduce transformation efficiency.
4. Pipet 1 µL of DNA into each micro centrifuge tube.
5. Pipette 50µl of competent cells into 1.5ml tube: 50µl in a 1.5ml tube per transformation. Tubes should be labeled, pre-chilled, and in a floating tube rack for support. Keep all tubes on ice. Don’t forget a 1.5ml tube for your control.
6. Pipette 1µl of resuspended DNA into 1.5ml tube: Pipette from well into appropriately labeled tube. Gently pipette up and down a few times. Keep all tubes on ice.
7. Pipette 1µl of control DNA into 2ml tube: Pipette 1µl of 10pg/µl control into your control transformation. Gently pipette up and down a few times. Keep all tubes on ice.
8. Close 1.5ml tubes, incubate on ice for 30min: Tubes may be gently agitated/flicked to mix solution, but return to ice immediately.
9. Heat shock tubes at 42°C for 45 sec: 1.5ml tubes should be in a floating foam tube rack. Place in water bath to ensure the bottoms of the tubes are submerged. Timing is critical.
10. Incubate on ice for 5min: Return transformation tubes to ice bucket.
11. Pipette 950µl SOC media to each transformation: SOC should be stored at 4°C, but can be warmed to room temperature before use. Check for contamination.
12. Incubate at 37°C for 1 hours, shaking at 220rpm
13. Pipette 100µL of each transformation onto petri plates Spread with sterilized spreader or glass beads immediately. This helps ensure that you will be able to pick out a single colony.

Day 3 : Cell growth, sampling, and assay

14. Set your instrument to read OD600 (as OD calibration setting)
15. Measure OD600 of the overnight cultures
16. Record data in your notebook
17. Import data into Excel ( Dilution Calculation) Sheet_1 provided
18. Dilute the cultures to a target OD 600 of 0.02 (see the volume of preloading culture and media in Excel ( Dilution Calculation ) Sheet_1) in 12 m l LB medium + Chloramphenicol
19. in 50 mL falcon tube (amber, or covered with foil to block light).
20. Incubate the cultures at 37°C and 220 rpm.
21. Take 500 μL samples of the cultures at 0, 2, 4, and 6 hours of incubation. (At each time point, you will take a sample from each of the 8 devices, two colonies per device, for a total of 16 samples per time point)
22. Place samples on ice.
23. At the end of sampling point you need to measure your samples (OD and Fl measurement), see the below for details.
24. Record data in your notebook
25. Import data into Excel ( cell measurement tab ) Sheet_1 provided

Measurement:

It is important that you use the same instrument settings that you used when measuring the fluorescein standard curve. This includes using the sample volume (100 ul) you used for the fluorescein measurement.

Samples should be laid out according to Fig. 2. Pipette 100 μl of each sample into each well. Replicate samples of colony #1 should be pipetted into wells in rows A, B, C and D. Replicate samples of colony #2 should be pipetted into wells in rows E, F, G and H. Set the instrument settings as those that gave the best results in your calibration curves (no measurements off scale). If necessary you can test more than one of the previously calibrated settings to get the best data (no measurements off scale).

Hint:

No measurement off scale means the data you get does not out of range of your calibration curve.

Layout for Abs600 and Fluorescence measurement:

At the end of the experiment, you should have four plates to read. Each plate should be set up as shown below. You will have one plate for each time point: 0, 2, 4, and 6 hours.