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Any deviation from these protocols will be noted in the Labnotes

Protocols

Colony PCR

CP1.1

How to amplify DNA from bacteria colonies and solutions

Important remarks

All solutions should be kept at ice until run of PCR

Materials
Premix Pfu-PCR

For 1 PCR reaction:

• 5 µL Pfu-buffer + MgSO4

• 1.5 µL 10mM dNTP mix

• 1.5 µL 10pmol/µL forward primer

• 1.5 µL 10pmol/µL reverse primer

• 0.5 µL Pfu polymerase enzyme (add just before PCR run)

• 40 µL H2O

Total vol.: 5O µL

Premix TAQ-PCR (no proofreading):
For 1 PCR reaction

• 2.5 µL 10x TAQ-buffer + MgCl2

• 0.5 µL 10mM dNTP mix

• 1.25 µL 10pmol/µL forward primer

• 1.25 µL 10pmol/µL reverse primer

• 19.25 µL Water

• 0.25 µL TAQ polymerase enzyme (add just before PCR run)

Total vol.: 25 µL

The TAQ polymerase has no proofreading, and should therefore only be used for size determination of DNA fragments. When PCR-product is to be purified and used for further experiments always use Phu polymerase!!! Make enough premix for your number of colonies +3.

Protocol
Colony PCR:

1. Select and transfer a single colony to a PCR tube with a pipette tip (afterwards use the same for plating out on plates)

2. Add all of the H2O used in the premix to the PCR tubes and place them in the microwave at full power for 2 min. with an open lid.

3. Make the premix (without water). Do not add enzyme until just before premix is added to the PCR tubes. Mix the premix by pipetting up and down (do not vortex!)

4. Add premix to each PCR tube.

5. Run PCR

PCR of DNA in solutions:

1. Transfer 1-2 µL of DNA to each PCR tube (to obtain the correct total volume adjust the volume of the H2O)

2. Make the premix(do not add enzyme until just before premix is added to the PCR tubes). Mix the premix by pipetting up and down (do not vortex!)

3. Add premix to each PCR tube.

4. Run PCR.

PCR program:

Other programs have been used as well

Pfu-PCR

1	Start	95°C	2 min
2	Denaturing	95°C	30 sec
3	Annealing	54°C	30 sec
4	Elongation	72°C	1 min
5	GOTO 2		rep. 29x
6	End	72°C	5 min
7	Hold	4°C

TAQ-PCR

1	Start	94°C	2 min
2	Denaturing	94°C	1 min
3	Annealing	55°C	1 min
4	Elongation	72°C	2min
5	GOTO 2		rep. 29x
6	End	72°C	5 min
7	Hold	4°C

CP1.2

Updated Taq protocol for length determination. Due to Taq's lack of proofreading, only use this protocol for length determination.

Protocol:
1. A single colony is transfered to each eppendorf tube with a pipette tip. (The same tip is used to plate out on a LA+antibiotic plate afterwards)
2. Add 30 ul H2O to each tube.
3. Microwave with open lid at full power for 2 minutes.
4. Prepare Pre-Mix (number of colonies+1) Distribute 19ul to each tube.
5. shortly spin down PCR tubes 6. Load and set PCR machine
7. Add TAQ polymerase at last moment. Make sure to get it under the surface of the solution.
8. Run PCR reaction.

Pre-mix:
5ul 10x TAQ buffer
2ul MgCl2 (Increase in 0.25ul incriments if the DNA you want to extract is longer than 3kb.)
2ul 10pmol/ul forward primer
2ul 10pmol/ul reverse primer
1ul 10mM dNTP mix
7ul H2O

1ul TAQ polymerase -> NB! Pre-Mix is made without TAQ polymerase!
PCR program

1	Start	94°C	2min
2	Denaturing	94°C	1min
3	Annealing	55°C	1min
4	Elongation	72°C	2min
5	GOTO 2		rep. 29x
6	End	72°C	3min
7	Hold	4°C	-

Elongation time is adjusted according to the length of the template. (1 min for every 1Kbp)

CP1.3

Updated Taq protocol for length determination. Due to Taq's lack of proofreading, only use this protocol for length determination.

Protocol:
1. A single colony is transfered to each eppendorf tube with a pipette tip. (The same tip is used to plate out on a LA+antibiotic plate afterwards)
2. Add 15 ul H2O to each tube.
3. Microwave with open lid at full power for 2 minutes.
4. Prepare Pre-Mix (number of colonies+1) Distribute 9.5ul to each tube.
5. shortly spin down PCR tubes 6. Load and set PCR machine
7. Add TAQ polymerase at last moment. Make sure to get it under the surface of the solution.
8. Run PCR reaction.

Pre-mix:
2.5ul 10x TAQ buffer
1ul MgCl2 (Increase in 0.25ul incriments if the DNA you want to extract is longer than 3kb.)
1ul 10pmol/ul forward primer
1ul 10pmol/ul reverse primer
0.5ul 10mM dNTP mix
3.5ul H2O

0.5ul TAQ polymerase -> NB! Pre-Mix is made without TAQ polymerase!
PCR program

1	Start	94°C	2min
2	Denaturing	94°C	1min
3	Annealing	55°C	1min
4	Elongation	72°C	2min
5	GOTO 2		rep. 29x
6	End	72°C	3min
7	Hold	4°C	-

Elongation time is adjusted according to the length of the template. (1 min for every 1Kbp)
--Tipi 07:40, 22 July 2010 (UTC)

Making competent cells of E. coli for transformation

CC1.1

How to make competent cells for transformation

Compotent cells enough for 12 transformations

Important remarks
Use 2 ml eppendorf tubes

Cool eppendorf tubes at 4°C prior to use

Cool 50 ml 50 mM CaCl2 at 4°C prior to use

Materials
• ON culture of TOP10 cells in LB media

• Ice cold 50mM CaCl2

• LB media (pre-heated to 37°C)

Protocol
1. Dilute the culture to OD550 = 0,02 in 110 ml of LB. Incubate at 37°C with shaking until OD550 reaches 0.5

2. Divide the cells in 2x55 ml and transfer to Falcon tubes (the can hold only 55 ml). From now on proceed with the 2 tubes in parallel

3. move the CaCl2 to -20°C

4. Harvest cells by centrifugation at 4100 rpm (2160 G) at 4°C for 10 min.

5. Discard the supernatant (keep the cells on ice!)

6. Resuspend cells gently in 5 ml ice cold CaCl2 (50 mM) taken from -20°C and kept on ice.

7. Repeat the centrifugation step.

8. Discard the supernatant and resuspend cells in 1.2 ml of icecold CaCl2 (keep the cells on ice!)

9. Leave the cells on ice for 30 min => now the cells are ready for transformation.

CC1.2

How to make competent cells for transformation – the iGEM way

Important remarks
All of the experiment needs to be carried out in the micro lab

Materials
• SOB media (see separate protocol)

• Ice cold CCMB80 buffer

10 mM KOAc pH 7.0 (10 ml of a 1M stock/L)

80 mM CaCl2 .2H2O (11.8g/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 0.1 M HCl if nessessary.

Adjusting pH down will precipitate manganese dioxide from Mn containing solutions.

Sterile filter and store at 4°C. Slight dark precipitate appears not to affect its function.

• Top10 cells grown on SOB plate

• Glycerol

• SOC (see separate protocol)

Protocol
Preparing seed stocks.

1. Streak Top10 cells on an SOB plate and grow for single colonies at 23°C. (room temperature)

2. Pick single colonies into 2 mL of SOB medium and shake overnight at 23°C.

3. Add glycerol to 15%

4. Aliquot 1mL samples to Nunc cryotubes

5. Place tubes into a zip lock bag, immerse bag into a dry ice/ethanol bath for 5 min. (this step may not be necessary)

6. Place in -80°C freezer indefinetly

Preparing competent cells
1. 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 h.) You can adjust this temperature somewhat to fit your schedule aim for lower, not higher OD if you cannot hit this mark.

2. Cebtrifuge at 3000g at 4°C for 10 min. in a flat bottom centrifuge bottle (flat bottom centrifuge tubes make the fragile cells easier to resuspend pellets by mixing before adding large amounts of buffer).

3. Gently resuspend in 80 mL of ice cold CCMB80 buffer. (sometimes this is less than completely gentle. It still works).

4. Incubate on ice for 20 min.

5. Centrifuge again at 4°C and resuspend in 10 mL of ice cold CCMB80 buffer.

6. Test OD of a mixture of 200 µL SOC and 50 µL of the resuspended cells.

7. Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test.

8. Incubate on ice for 20 min.

9. Aliquot to chilled screw top 2 mL vials or 50 µL into chilled microtiter plates.

10. Store at -80°C indefinitely.

11. Optional: Test competence.

Measurement of competence

How to test transformation efficiency of competent cells – the iGEM way

Materials
• pUC19 plasmid (Invitrogen)

• SOC medium (see separate protocol)

Protocol
1. Transform 50 µL of competent cells with 1 µL of standard pUC19 plasmid. This is at 10 pg/µL or 10-5 µg/µL (This can be done by diluting 1 µL of NEB pUC19 plasmid (1 µg/µL, NEB part number NS3401S) into 100 mL of TE))

2. Keep on ice for 30 min.

3. Heat shock 60 s. at 42°C (Very important)

4. Add 250 µL SOC medium

5. Incubate at 37°C for 1 h. in 2 mL centrifuge tubes. (these tubes works well with transformation)
- Transformation with plasmids pSB1AC3 and pSB1AT3, which are chloramphenicol and tetracycline resistant, incubating for 2 h. yields many more colonies.

6. Plate 20 µL on AMP plates and spread.

7. Incabate plates at 37°C over night.

8. Count the colonies (good cells should yield around 100-400 colonies) Transformation efficiency is (dilution factor = 15) x colony x 105/µg DNA

We expect that the transformation efficiency should be between 5x108 and 5x109 cfu/µgDNA

Transformation

TR1.1

How to transform compotent cells
Important remarks
Pre-heat LB media to 37°C

Pre-dry LA plates with the appropriate antibiotics.

Pre-cool 2 mL eppendorf tubes.

Keep cells on ice at all times!!

Remember controls:
Positive control with your uncut vector

Negative control with no DNA

Materials
• Freshly made compotent E. coli cells.

• LA plates with appropriate antibiotics

• LB media

Protocol
1. Transfer 5 µl DNA (plasmid or ligation mix) to precooled eppendorf tubes. (Use only 1ul if DNA is taken from distribution plates.)

2. Transfer 200 µl of cells with to the tube and mix by pipetting up and down (keep the cells on ice at all times)

3. Leave on ice for 30 min.

4. Heat shock for 90 sec. at 42°C in a water bath, do not shake tubes.

5. Place on ice for 2 min.

6. Add 1.5 mL of preheated LB media (37°C)

7. Incubate at 37°C for 1 hour with gentle shaking.

8. Plate 2 plates with 150 µl mixture on LA plates with the appropriate antibiotics.

9. Pellet the remaining cells 5 min at 3500 rpm and discard approximately 900 µl of the supernatant.

10. Resuspend cells and plate out on LA plates with appropriate antibiotics.

11. Incubate all plates ON at 37°C

Restriction digest

RD1.1

How to digest DNA using fast digest restriction enzymes.

Important remarks
Remember to load a documentation slot next to the marker and take a picture of this for later documentation.

Materials
Restriction mixture:

For 1 digest reaction.

• 12 µL H2O (or 13 µL if only one restriction enzyme is used)

• 1 µL enzyme A

• 1 µL enzyme B

• 2 µL Fast Digest green buffer

• 5 µL PCR product

Multiply protocol if more digested PCR product is needed

Protocols
1. Prepare a purification gel

2. Mix the restriction mixture in en eppendorf tube by pipetting up and down

3. Leave for 5 min. at 37°C (no shaking!)

4. Immidiately load the restriction mixture in the gel

5. Run the gel and perform a purification step

Ligation

LG1.1

How to assemble DNA biobricks

Materials
Ligation mixture:

For 1 ligation reaction

• 2 µL 10x T4 ligase buffer

• 1 µL T4 ligase (add last!)

• 5 µL PCR product (cut) of each brick which is to be ligated – or 1 part plasmid and 5 part bricks

Protocol
1. Prepare the ligation mixture and mix by pipetting up and down

2. Leave the mixture over-night at 17°C

3. Test ligation using TAQ-PCR and run test gel afterwards in order to check that the PCR product has the right size

LG1.2

How to assemble DNA biobricks

Materials
Ligation mixture:

For 1 ligation reaction

• 2 µL 10x T4 ligase buffer

• 1 µL T4 ligase (add last!)

• 5 µL PCR product (cut) of each brick which is to be ligated – or 1 part plasmid and 5 part bricks

• Add H2O to reach a total volume of 20mL

Protocol
1. Prepare the ligation mixture and mix by pipetting up and down

2. Leave the mixture over-night at 17°C

3. Test ligation using TAQ-PCR and run test gel afterwards in order to check that the PCR product has the right size

--Tipi 06:48, 20 July 2010 (UTC)

LG1.3

Materials

• T4 DNA ligase
• 10x T4 DNA Ligase Buffer
• Deionized, sterile H₂O
• Purified, linearized vector (likely in H₂O or EB)
• Purified, linearized insert (likely in H₂O or EB)

Equipment Vortex Procedure 10μL Ligation Mix Larger ligation mixes are also commonly used

• 1.0 μL 10X T4 ligase buffer
• 6:1 molar ratio of insert to vector (~10ng vector)
• Add (8.5 - vector and insert volume)μl ddH₂O
• 0.5 μL T4 Ligase

Calculating Insert Amount <math>{\rm Insert\ Mass\ in\ ng} = 6\times\left[\fracTemplate:\rm Insert\ Length\ in\ bpTemplate:\rm Vector\ Length\ in\ bp\right]\times{\rm Vector\ Mass\ in\ ng}</math> The insert to vector molar ratio can have a significant effect on the outcome of a ligation and subsequent transformation step. Molar ratios can vary from a 1:1 insert to vector molar ratio to 10:1. It may be necessary to try several ratios in parallel for best results. Method

1. Add appropriate amount of deionized H₂O to sterile 0.6 mL tube
2. Add 1 μL ligation buffer to the tube.
   Vortex buffer before pipetting to ensure that it is well-mixed.
   Remember that the buffer contains ATP so repeated freeze, thaw cycles can degrade the ATP thereby decreasing the efficiency of ligation.
3. Add appropriate amount of insert to the tube.
4. Add appropriate amount of vector to the tube.
5. Add 0.5 μL ligase.
   Vortex ligase before pipetting to ensure that it is well-mixed.
   Also, the ligase, like most enzymes, is in some percentage of glycerol which tends to stick to the sides of your tip. To ensure you add only 0.5 μL, just touch your tip to the surface of the liquid when pipetting.
6. Let the 10 μL solution sit at 22.5°C for 30 mins
7. Denature the ligase at 65°C for 10min
8. Dialyze for 20 minutes if electroporating
9. Use disks shiny side up
10. Store at -20°C

Factors affecting efficiency From Tom Ellis A protocol analysis experiment for a typical DNA ligation (7.2 kb vector + 0.6 kb insert, sticky ends) gave optimal ligation efficiency when 50 ng of vector was ligated overnight at 16°C with a 2:1 insert:vector molar ratio and standard T4 ligase. Ligase was heat inactivated at 65°C for 20 mins before 2 μL (of 20 μL) was used to transform commercial heat-shock competent cells. Ligation efficiency was marginally decreased by

1. Doing a 1 hr ligation at room temperature
2. Using 100 ng vector
3. Using insert:vector molar ratios of 5:1 and 1:1

Ligation efficiency was noticably decreased (x100) by

1. Sticky end ligation with a larger insert (5.2 kb vector + 2.6 kb insert)
2. Blunt end ligation

Ligation efficiency was severely decreased (x10000) by

1. Using DNA fragments that have been exposed to UV during the gel extraction procedure (can avoid by blind excision, or by using a black-light or 365nm UV transilluminator instead of the usual 312nm type)
2. Using the NEB Quick Ligation Kit (heat inactivation of PEG in the buffer ruins transformation, without heat inactivation the ligation probably would've been fine)

For additional troublshooting, check out the NEB FAQ page for T4 ligation: [1] Notes

1. Make sure the buffer is completely melted and dissolved. The white precipitate is BSA according to NEB. Make sure the buffer still smells strongly like "wet dog" (to check if the DTT is still good).
2. Because ligase buffer contains ATP, which is unstable and degraded by multiple freeze/thaw cycles, you may want to make 10-20ul aliquots from the original tube. Ligase buffer may be spiked with additional ATP.
3. If you are having trouble with your ligation, NEB offers FAQ's (Quick Ligation T4 DNA ligase) and tips to help.
4. Prior to the ligation, some heat their DNA slightly (maybe ~37°C) to melt any sticky ends which may have annealed improperly at low temperatures.
5. Tom Knight has read that ligase can inhibit transformation Michelsen-Anal-1995. By heat-inactivating the ligase, this inhibition can be avoided. However, according to the NEB FAQ, heat-inactivation of PEG (which is present in the ligation reaction) also inhibits transformation, therefore a spin-column purification is recommended prior to transformation if you are having problems.
6. Treating PCR products with proteinase K prior to restriction digest dramatically improves the efficiency of subsequent ligation reactions. Crowe-NAR-1991
7. Using SYBR Safe DNA Gel Stain is a safer, non-carcinogenic alternative to ethidium bromide.
8. T4 DNA Ligase is very sensitive to shear, so spinning your ligation mix or vortexing it to mix it can affect your yields. Instead try mixing with the pipette tip or slowly resuspending the solution.

Acknowledgments This protocol is primarily based on Endy:DNA ligation using T4 DNA ligase. References <biblio>

1. Crowe-NAR-1991 pmid=2011503
2. Olivera-PNAS-1967 pmid=5341238

// DNA ligation by Escherichia coli DNA ligase

1. Michelsen-Anal-1995 pmid=7778774

</biblio>

DNA extraction from gel (fermentas)

DE1.1

How to extract and purify DNA from gel

Important remarks
All steps should be carried out at room temperature.

All centrifugations should be carried out in a table-top microcentrifuge at >12000x g

Materials
• Binding buffer

• Wash buffer (diluted with ethanol)

• Elution buffer

Protocol
1. Weigh a 1.5 µL tube

2. Excise gel slice containing the DNA fragment using a clean scalpel (cut as close to the DNA as possible)

3. Place the gel slice into the pre-weighed tube and record the weight of the gel slice

4. Add 1:1 volume of binding buffer to the gel slice (e.g. add 100 µL of binding buffer for every 100 mg of agarose gel)

5. Incubate the gel mixture at 50-60°C for 10 min. or until the gel slice is completely dissolved. Mix the tube by inversion every few minutes. The color of the solution should be yellow. If the color of the solution is orange or violet add 10 µL 3M sodium acetate , pH 5.2 and mix. The color will then turn yellow.

6. Transfer up to 800 µL of the solubilized gel solution to the GeneJET purification column. Centrifuge for 1 min. Discard the flow-through and place column back into the same collection tube.

7. Add 700 µL of Wash buffer to the column. Centrifuge for 1 min. Discard flow-through and place the column back into the collection tube.

8. Centrifuge the empty column for an additional 1 min. to completely remove residual Wash buffer This step is essential to avoid residual ethanol in the purified DNA solution

9. Transfer the column into a clean 1.5 mL microcentrifuge tube. Add 50 µL of Elution buffer to the center of the column membrane. Centrifuge for 1 min.

10. Discard the column and store the purified DNA at -20°C.

DE1.2

How to extract and purify DNA from gel

Important remarks
All steps should be carried out at room temperature.

All centrifugations should be carried out in a table-top microcentrifuge at >12000x g

Materials
• Binding buffer

• Wash buffer (diluted with ethanol)

• Elution buffer

Protocol
1. Weigh a 1.5 µL tube

2. Excise gel slice containing the DNA fragment using a clean scalpel (cut as close to the DNA as possible)

3. Place the gel slice into the pre-weighed tube and record the weight of the gel slice

4. Add 1:1 volume of binding buffer to the gel slice (e.g. add 100 µL of binding buffer for every 100 mg of agarose gel)

5. Incubate the gel mixture at 50-60°C for 10 min. or until the gel slice is completely dissolved. Mix the tube by inversion every few minutes. The color of the solution should be yellow. If the color of the solution is orange or violet add 10 µL 3M sodium acetate , pH 5.2 and mix. The color will then turn yellow.

6. Transfer up to 800 µL of the solubilized gel solution to the GeneJET purification column. Centrifuge for 1 min. Discard the flow-through and place column back into the same collection tube.

7. Add 700 µL of Wash buffer to the column. Centrifuge for 1 min. Discard flow-through and place the column back into the collection tube.

8. Centrifuge the empty column for an additional 1 min. to completely remove residual Wash buffer This step is essential to avoid residual ethanol in the purified DNA solution

9. Transfer the column into a clean 1.5 mL microcentrifuge tube. Add 20 µL of H2O to the center of the column membrane. Centrifuge for 1 min.

10. Discard the column and store the purified DNA at -20°C.

--Tipi 06:45, 20 July 2010 (UTC)

DE1.3

Protocol for purification of DNA from TAE and TBE agarose gel bands
Kit from GFX.

Sampla capture
1. Weigh a DNase-free 1.5 ml microcentrifuge tube.

2. Exice band of interest from the gel and place in microcentrifuge tube.

3. Weigh microcentrifuge tube plus agarose gel band.

4. Calculate weight of agarose gel slice.

5. Add 10 ul Capture buffer type 3 for each 10 mg agarose slive (add at least 300 ul!)

6. Mix by inversion.

7. Place at 60 degrees celcius until agarose is completely dissolved.

Sample binding
1. Add up to 600 ul Capture buffer-sample mix to assembled GFX MicroSpin columns and Collection tube.

2. Leave at room temperature for 1 minute.

3. Centrifuge for 30 sec. at 16,000g.

4. Discard the flow-through in the Collection tube and place the MicroSpin column in the collection tube again.

5. Repeat sample binding step until all sample is loaded onto the MicroSpin column.

Wash and Dry
1. Add 500 ul Wash buffer type 1.

2. Centrifuge for 30 sec. at 16,000g.

3. Discard flow-through and keep Collection tube a above.

4. Centrifuge again for 30 sec. at 16,000g.
More flow-through will appear in the collection tube. It is important to centrifuge this second time to get the sample completely dry. This step is not included in the original protocol.

5. Discard Collection tube and trensfer MicroSpin column to a clean 1.5 ml DNase-free microcentrifuge tube.

Elution
1. Add 10 - 50 ul Elution buffer type 4 or 6. (10 ul is fine for small volumes)

2. Leave at room temperature for 60 sec.

3. Centrifuge for 1 min. at 16,000g.

4. Retain flow-through and discard MicroSpin Column.

5. Store purified sample DNA at -20 degrees or proceed to cutting DNA og ligation.

Genomic DNA purification

GP1.1

How to extract and purify genomic DNA

Important remarks
All steps should be carried out at room temperature.

Be sure to mix thoroughly when adding the solutions.

Addition and removal of chloroform should be carried out in fume hood.

Materials
• Lysis solution

• Chloroform

• Precipitation solution (80 µL is diluted in 720 µL of H2O just prior to use)

• 1.2M NaCl solution

• Ice cold ethanol (70%)

• H2O

Protocol
1. Mix 200 µL of sample (ON culture) with 400 µL of Lysis solution and incubate at 65°C for 5 min.If a frozen sample is used lysis solution should be added before thawing and incubated at 65°Cfor 10 min. with occasional inverting the tube.

2. Immediately add 600 µL of chloroform, gently emulsify by inversion (3-5 times) and centrifuge the sample at 10.000 rpm for 2 min.

3. Prepare precipitation solution.

4. Transfer the upper aqueous phase containing DNA to a new tube and add 800 µL of freshly prepared precipitation solution, mix gently by several inversions at room temperature for 1-2 min. and centrifuge at 10.000 rpm for 2 min.

5. Remove supernatant completely (do not dry) and dissolve DNA pellet in 100 µL of 1.2M NaCl solution by gentle vortexing (make sure that the pellet is completely dissolved) To avoid loosening the pellet, keep the tube in the same angle as when placed in the centrifuge!

6. Add 300 µL of cold ethanol, let the DNA precipitate (10 min. at -20°C) and spin down (10.00 rpm, 3-4 min.).Pour off the ethanol and dissolve DNA in 15 µL of sterile dH2O by gentle vortexing.

7. Measure DNA concentration on nanodrop.

8. Store DNA at -20°C

Plasmid miniprep kit (Fermentas)

MP1.1

How to isolate plasmids from cultures

Important remarks
All steps should be carried out at room temperature.

Step 1 and 2 must be carried out in the micro lab.

Materials
• Resuspension solution (with RNase A)

• Lysis solution

• Neutralization solution

• Wash solution (diluted with ethanol)

• Elution solution

Protocol
1. Resuspend pelleted cells in 250 µL Resuspension solution. Resuspend completely by vortexing. Transfer the cell suspension to microcentrifuge tubes.

2. Add 250 µL Lysis solution and mix thoroughly by inverting the tube 4-6 times until the solution is viscous and slighty clear. (Do not vortex!)

3. Add 350 µL Neutralization buffer and mix immediately and thoroughly by inverting the tube 4-6 times. (It is important to mix gently to avoid localized precipitation)

4. Centrifuge for 5 min. to pellet cell debris and chromosomal DNA.

5. Transfer supernatant to the supplied GeneJet spin column, without disturbing or transferring the white precipitate.

6. Cenntrifuge for 1 min. Discard flow-through and place column back into the same collection tube.

7. Add 500 µL Wash solution to the column. Centrifuge for 30-60 s. and discard the flow-through. Place column back into the same tube.

8. Repeat step 7.

9. Discard the flow-through and centrifuge for an additional 1 min. to remove residual wash solution. (This step is essential to avoid residual ethanol in plasmid preps)

10. Transfer the column into a fresh 1.5 mL microcentrifuge tube. Add 50 µL of Elution buffer to the center of the column membrane to elute the plasmid DNA (do not touch the membrane with the pipette tip!). Incubate for 2 min. at room temperature and centrifuge for 2 min.
Optional: repeat elution step to increase the overall yield by 10-20%.

11. Discard the column and store the purified plasmid DNA at -20°C.

MP1.2

How to isolate plasmids from cultures

Important remarks
All steps should be carried out at room temperature.

Step 1 and 2 must be carried out in the micro lab.

Materials
• Resuspension solution (with RNase A)

• Lysis solution

• Neutralization solution

• Wash solution (diluted with ethanol)

• Elution solution

Protocol
1. Transfer 10 mL ON-culture to a 15 mL falcon tube and spin down at 4000g for 15 min.

2. Resuspend pelleted cells in 500 µL Resuspension solution. Resuspend completely by vortexing. Divide the cell suspension in 2x250ul and transfer to eppendorf tubes. From now on proceed with the two tubes in parallel.

3. Add 250 µL Lysis solution and mix thoroughly by inverting the tube 4-6 times until the solution is viscous and slighty clear. (Do not vortex!)

4. Add 350 µL Neutralization buffer and mix immediately and thoroughly by inverting the tube 4-6 times. (It is important to mix gently to avoid localized precipitation)

5. Centrifuge for 5 min. to pellet cell debris and chromosomal DNA.

6. Transfer supernatant to the supplied GeneJet spin column, without disturbing or transferring the white precipitate.

7. Cenntrifuge for 1 min. Discard flow-through and place column back into the same collection tube.

8. Add 500 µL Wash solution to the column. Centrifuge for 30-60 s. and discard the flow-through. Place column back into the same tube.

9. Repeat step 8.

10. Discard the flow-through and centrifuge for an additional 1 min. to remove residual wash solution. (This step is essential to avoid residual ethanol in plasmid preps)

11. Transfer the column into a fresh 1.5 mL microcentrifuge tube. Add 50 µL of Elution buffer to the center of the column membrane to elute the plasmid DNA (do not touch the membrane with the pipette tip!). Incubate for 2 min. at room temperature and centrifuge for 2 min.
Optional: repeat elution step to increase the overall yield by 10-20%.

12. Discard the column and store the purified plasmid DNA at -20°C.

MP1.3

How to isolate plasmids from cultures

Important remarks
All steps should be carried out at room temperature.

Step 1 and 2 must be carried out in the micro lab.

Materials
• Resuspension solution (with RNase A)

• Lysis solution

• Neutralization solution

• Wash solution (diluted with ethanol)

• Elution solution

Protocol
1. Cells from ON culture is reboosted by transfering 2.5mL ON culture to 15mL LB preheated LB medium.Cells are then grown for additionally 2 hours.

2. Transfer all 17.5mL new culture to a 50mL falcon tube10 mL ON-culture and spin down at 4000g for 15 min.

3. Resuspend pelleted cells in 250 µL Resuspension solution. Resuspend completely by vortexing.

4. Add 250 µL Lysis solution and mix thoroughly by inverting the tube 4-6 times until the solution is viscous and slighty clear. (Do not vortex!)

5. Add 350 µL Neutralization buffer and mix immediately and thoroughly by inverting the tube 4-6 times. (It is important to mix gently to avoid localized precipitation)

6. Centrifuge for 5 min. to pellet cell debris and chromosomal DNA.

7. Transfer supernatant to the supplied GeneJet spin column, without disturbing or transferring the white precipitate.

8. Cenntrifuge for 1 min. Discard flow-through and place column back into the same collection tube.

9. Add 500 µL Wash solution to the column. Centrifuge for 30-60 s. and discard the flow-through. Place column back into the same tube.

10. Repeat step 8.

11. Discard the flow-through and centrifuge for an additional 1 min. to remove residual wash solution. (This step is essential to avoid residual ethanol in plasmid preps)

12. Transfer the column into a fresh 1.5 mL microcentrifuge tube. Add 50 µL of Elution buffer to the center of the column membrane to elute the plasmid DNA (do not touch the membrane with the pipette tip!). Incubate for 2 min. at room temperature and centrifuge for 2 min.
Optional: repeat elution step to increase the overall yield by 10-20%.

13. Discard the column and store the purified plasmid DNA at -20°C.

Preparation of Agarose for gel electrophoresis

AG1.1

How to prepare Agarose for gel electrophoresis.

Important remarks
Agarose concentration is dependent on the size of the DNA fragment that needs to be seperated (see the door of the incubator in the gel room)

Addition of EtBr is carried out in fume hood.

Materials
• Seachem Agarose

• TAE buffer

• EtBr

Protocol
1. For a 1% agarose gel mix 3 g agarose and 300 mL TAE buffer in a 500 mL flask.

2. The mixture is heated for 5 min. at max temperature in micro-wave. Remember to note name, date and –EtBr on the flask.

3. Place flask in the incubator for 20 min or at room temperature until cooled to 60°C.

4. Add 5 droplets of EtBr.

5. Cast gel and leave for 20 min until the gel is set. Remaining agarose solution is placed in incubater for later use.

6. Load gel and run gel. Load only 5 µL of DNA marker

Preparation of SOB and SOC media

How to prepare SOB and SOC media for transformation.

SOB medium

Used in growing bacteria for preparing chemically compotent cells.

Materials
For 1 L:
• 20 g tryptone

• 5 g yeast extract

• 0.5g NaCl

• dH2O to 1 L

• KCl (is made by dissolving 1.86 g of KCl in 100 mL of deionized H2O)

• 2M MgCl2 (is made by dissolving 19g MgCl2 in 90 mL dH2O =>adjust to obtain a volume of 100 mL using dH2O => autoclavate)

Protocol
1. Add tryptone, yeast extract and NaCl to 950 mL of dH2O and shake until solute has dissolved.

2. Add 10 mL of 250 mM solution of KCl

3. Adjust volume to 1 L using dH2O

4. Autoclavate for 20 min.

5. Just before use add 5 mL of sterile solution of 2M MgCl2

SOC medium

Materials
• SOB medium.

• 1M glucose (is made by dissolving 18g of glucose in 90 mL of dH2O => adjust to obtain a volume of 100 mL using dH2O)

Protocol
1. Cool SOB medium to 60°C

2. Add 20mL of 1M glucose.

Extraction af Carotenoids and polyene chromophores

EX1.1

1. Incubate E. Coli in 100 ml LB media with appropriate antibiotics at 37 oC for 20 hours
2. Harvest cells using centrifugation at 4000 G for 15 min
3. Re-suspend cells in 8 mL acetone and sonicate the sample for 2x 2 min
4. Centrifuge the samples at 16000 G for 2 min and collect the supernatant
5. Measure absorbance using UV-Vis spectrophotometer at 450 nm, was preformed on a …. From MEMPHYS

EX1.2

1. Incubate E. Coli in 100 ml LB media with appropriate antibiotics at 37 oC for 20 hours
2. Harvest cells using centrifugation at 4000 G for 15 min
3. Re-suspend cells in 8 mL acetone and sonicate the sample for 2x 2 min
4. Centrifuge the samples at 16000 G for 2 min and collect the supernatant
5. Measure absorbance using an HPLC at 450 nm using a C18… column with 50% Acetonitrill, 50% water and 10% Acetonitrill 90% Methanol as eluents A and B

Photosensor characterisation

PS1.1

Materials
• Diluted LB media
• Difco Agar
• 1mM retinal

Swimming motility plates
1. LB media is mixed with 0.3% difco agar and is autoclavated
2. The appropriate antibiotic and 1uM retinal is added to the autoclaved media (NB: for the media used for the control plates, no antibiotic or retinal is added)
3. Plates are cast and incubated ON at room temperature
4. 15 minutes prior to the experiment the plates are incubated at 37°C.

Sample preparation
1. Colony is inoculated in 5mL LB media containing the appropriate antibiotic. The culture is grown ON at 37°C and 180rpm.
2. ON culture is diluted in 5mL LB media containing the appropriate antibiotic to reach an OD550 of 0.02 and are incubated at 37°C and 180rpm until it reaches an OD550 of 0.5.
3. 1mM retinal is added to the culture (NB: no retinal is added to the cultures containing the control cells)
4. The tubes containing the cultures are wrapped in tin foil and are subsequently grown for 2 hours at 37°C and 180rpm.

Motility assay
1. 2x2.5uL of culture is placed on each plate, and the plates are placed in a specially engineered lightbox, so that ½ of each plate is illuminated with blue light and the other ½ is kept in dark. 1mL of each culture is used for microscopy.
2. The plates are incubated at 37°C for 24 hours.
3. Pictures are taken

Microscopy
1. 5uL cell culture is used for the microscopy
2. To avoid laminar flow, the microscopy slide is sealed with nail polish.
3. Samples are examined under the microscope.

Growth rate assay

GA1.1

Materials
• LB media
• Spectrophotometer

Protocol
1. A colony is inoculated in 5mL LB media containing the appropriate antibiotic.
2. The culture is incubated over night at 37°C and 180rpm.
3. The optical density at 550nm (OD550) of the ON culture is measured and the culture is diluted in 25uL fresh LB media containing the appropriate antibiotic to reach an OD550 of 0.02. The culture is incubated for 24 hours at 37°C and 180rpm.
4. OD550 of the colony is measured every hour for the first 12 hours, and after 24 hours.

Flagella staining

1.1

Day 1: The bacteria were grown in 5 ml-LB media ON. The solutions used for staining were prepared.

Solution I: The following components were added in the listed order:
• 5 g of tannic acid was dissolved in 9.65 ml distilled water.
• 150 µl 9% FeCl3
• 100 µl 1% NaOH
• 200 µl formalin

Solution II:
• 2 g silver nitrate was dissolved in 10 ml distilled water
• 10% aqueous ammonia solution was added until the silver nitrate was dissolved. Approximately 2 ml.

Day 2: The bacteria were boosted in 5 ml LB-media to ensure that they were in the exponential growth phase when used for staining. They were diluted to approximately OD550 1.

Preliminary bacteria work: • The bacteria were centrifuged 15 min at 4000rpm
• The pellet was resuspended in LB-media to an OD550 of 3.

Staining protocol:
• A clean glass slide was used and Poly-L-Lysin was added onto a small area.
• 20 µl of the bacteria solution was plated on the slide and allowed to air dry.
• The slide was flooded with solution I and allowed to stand for 30 min before it was washed with distilled water.
• Solution II was added and was incubated at room temperature for 10 min and was washed with distilled water.
• The slide was flooded with a carbol-fuchsin solution and air-dried before washed with distilled water.

PBS were added to the area containing the bacteria and they were covered with a cover slide. The slides are now ready for examination under the microscope.

1.2

Day 1: Bacteria were platede on agar plates and incubated at 37 degrees ON. The staining solutions were prepared.

Solution I:
The following components were added in the listed order:
• 5 g of tannic acid was dissolved in 9.65 ml distilled water.
• 150 µl 9% FeCl3
• 100 µl 1% NaOH
• 200 µl formalin

Solution II:
• 2 g silver nitrate was dissolved in 10 ml distilled water.
• 10% aqueous ammonia solution was added until the silver nitrate was dissolved. Approximately 2 ml.

Day 2: A bacteria colony was dissolved in LB-media.

Staining protocol:
• A clean glass slide was used and Poly-L-Lysin was added onto a small area.
• 20 µl of the bacteria solution was plated on the slide and allowed to air dry.
• The slide was flooded with solution I and allowed to stand for 30 min before it was washed with distilled water.
• Solution II was added and was incubated at room temperature for 10 min and was washed with distilled water.
• The slide was flooded with a carbol-fuchsin solution and air-dried before washed with distilled water.

PBS were added to the area containing the bacteria and they were covered with a cover slide. The slides are now ready for examination under the microscope.

Stability assay

SA 1.1

Stability assay

Materials

• LB media
• LA plates
• LA plates with appropriate antibiotic
• 0.9% NaCl

Protocol

1. A bacteria colony is inoculated in 5mL LB media with appropriate antibiotic.
2. Culture is incubated over night at 30°C and 180rpm
3. 100uL of the culture is serial diluted in 900uL 0.9% NaCl to reach a dilution of 107.
4. 100uL of the 105, 106 and 107 dilution is spreaded on LA plates, and LA plates with the appropriate antibiotic, respectively.
5. Plates are incubated at 37°C for 16-24 hours. The following day the colonies formed on the plates are counted and cfu are determined.
6. 500uL of the 105 dilution is added to 4.5mL of fresh LB media without any antibiotics.
7. The new culture is incubated over night at 30°C and 180rpm
8. The experiment is carried out for 5 days. (NB: antibiotic is only added to the first culture. The remaining days the bacteria are grown in cultures without any antibiotics)

Almost like cake recipes... although cake is infinitely more delicious than bacteria.

Contents 1 Protocols 1.1 Colony PCR 1.1.1 CP1.1 1.1.2 CP1.2 1.1.3 CP1.3 1.2 Making competent cells of E. coli for transformation 1.2.1 CC1.1 1.2.2 CC1.2 1.2.3 Measurement of competence 1.3 Transformation 1.3.1 TR1.1 1.4 Restriction digest 1.4.1 RD1.1 1.5 Ligation 1.5.1 LG1.1 1.5.2 LG1.2 1.5.3 LG1.3 1.6 DNA extraction from gel (fermentas) 1.6.1 DE1.1 1.6.2 DE1.2 1.6.3 DE1.3 1.7 Genomic DNA purification 1.7.1 GP1.1 1.8 Plasmid miniprep kit (Fermentas) 1.8.1 MP1.1 1.8.2 MP1.2 1.8.3 MP1.3 1.9 Preparation of Agarose for gel electrophoresis 1.9.1 AG1.1 1.10 Preparation of SOB and SOC media 1.10.1 SOB medium 1.10.2 SOC medium 1.11 Extraction af Carotenoids and polyene chromophores 1.11.1 EX1.1 1.11.2 EX1.2 1.12 Photosensor characterisation 1.12.1 PS1.1 1.13 Growth rate assay 1.13.1 GA1.1 1.14 Flagella staining 1.14.1 1.1 1.14.2 1.2 1.15 Stability assay 1.15.1 SA 1.1