Overview

1.Experiments description

1.1.Preparation of LB Broth

To begin the cloning experiment, we first prepared LB Broth. This required mixing 25 grams of LB Broth powder (Scharlau, cat# 02-385) with every 1 litre of distilled water. After thorough mixing, we autoclaved the solution at 121°C for 15 minutes, using a total volume of 0.5 litres. Subsequently, we completed the mixture by adding autoclaved water to reach the 1-litre mark.

Fig.1-LB broth liquid medium

1.2.Preparation of LB Agar

LB Agar was prepared by dissolving 20 grams of LB Agar powder (Scharlau, cat# 01-385) in 0.5 litres of distilled water. After heating the mixture to melt the agar, it was autoclaved at 121°C for 15 minutes. 25 μl of the agar solution were then poured onto each agar plate, and these plates were allowed to cool down.

1.3.Preparation of 1 M CaCl2 Solution

Our next step involved preparing a 1 M CaCl2 solution. Using CaCl2 anhydrous (C1016) with a molecular weight of 110.98, we calculated that 200 millilitres of the solution required approximately 22.2 grams of CaCl2. We mixed the calculated amount of CaCl2 with distilled water to obtain the desired 1 M CaCl2 solution.

1.4.Preparation of Competent Cells

On August 24, 2023, we initiated the preparation of competent E. coli cells for transformation. We began by plating 200 microliters, sourced from a glycerol stock, onto two plates, designating them as "DH5 alpha 1" and "DH5 alpha 2." These plates were specifically chosen for their capacity to support nearly confluent cell growth. The plates were subsequently stored at 4°C on August 25, 2023. On August 27, 2023, we performed colony selection by picking three colonies from each plate and placing them into 2 ml glass tubes. These tubes were then subjected to overnight shaking at 320 RPM and 37°C.

Fig.2- E.coli colonies on agar plate

On August 28, 2023, we transferred 250 μl from the overnight culture into 50 ml flasks, aiming to grow the culture until the optical density at 600 nm (O.D.600) reached the desired range of 0.4 - 0.6. Utilising a densitometer for monitoring, we noted that the overnight culture had a reading of 1.4. After 2:45 hours after the start of the growth, the density stood at 3, while at 3:30 hours, it had reached 5.1, leading us to halt the growth process.

Following growth, we centrifuged the cells at 5000 g for 5 minutes to obtain pellets. The pellets were resuspended in 25 ml of 0.1 M CaCl2, prepared using 2.5 ml of 1 M CaCl2. After an hour of incubation on ice, with a brief benchtop rest for 5 minutes, the suspension was subjected to a second centrifugation step at 5000 g for 5 minutes. Following the second centrifugation, the pellet was resuspended in 3 ml of a chilled solution consisting of 15% Glycerol and 0.1 M CaCl2. This solution was prepared by mixing 450 μl of glycerol, 300 μl of 1 M CaCl2, and 2.25 ml of H2O. Finally, we aliquoted 100 μl of the suspension and stored it at -80°C for future use.

1.5.Transformation of DH5 alpha Competent Bacteria

In the transformation process, competent bacteria were stored at -80°C in 100 microliter aliquots. For each transformation, 50 μl aliquots of competent cells were used. 1 μl of each plasmid (pSB1C30 and pJUMP26-1A) was transformed.

The cells were initially incubated on ice for 30 minutes. A heat shock was then performed, involving a brief exposure to 42°C for 40 seconds, followed by immediate placement back on ice for 2 minutes. Subsequently, 950 μl of LB medium were added to the cells. The cells were then shaken at 37°C for 1 hour. To complete the transformation process, 250 μl of the transformation solution were plated on the plates with the corresponding antibiotic (chloramphenicol for pSB1C30 and kanamycin for pJUMP26-1A).

1.6.Transformation of Plasmids and Colony Selection

We noted that the transformation of plasmids (pSB1C30 and pJUMP26-1A) resulted in colonies appearing only on the pJUMP26-1A plates. The following day, all bacterial colonies from the transformed plates were spun down and plated using 250 microliters of the bacterial culture. Colonies were observed on both types of plates this time, with pSB1C30 colonies notably displaying a distinctive red coloration, which confirmed that the transformation had succeeded, as pSB1C30 contains a red fluorescent protein. The pJUMP26-1A colonies had a green tint, which could also be thanks to the green fluorescent protein that the plasmid contains. Three selected colonies from both types of plates were grown overnight at 320 RPM and 37°C.

(a)

(b)

Fig.3 Transformation of plasmid DNA for pSB1C30(a) and pJUMP26-1A(b)

1.7.DNA Extraction Using QIAprep Miniprep

The DNA extraction process involved several steps to obtain purified DNA samples for further analysis.The bacterial cells were initially spun down at 5400 g for 10 minutes at 4°C and the pellet was then resuspended in 250 μl of Buffer P1. Subsequent steps involved the addition of Buffer P2, Buffer N3, centrifugation, and washing to remove impurities and elute the DNA. We added 250 μl of Buffer P2 and mixed thoroughly by inverting the tube 4–6 times. Then we added 350 μl of Buffer N3, mixing by inverting the tube 4–6 times. We then centrifuged for 10 minutes at 13000 rpm (~17,900 x g) in a table-top microcentrifuge and applied 800 μl of the supernatant to the QIAprep 2.0 Spin Column by pipetting. All the following centrifugations were done at 13000 rpm. Next we centrifuged for 1 minute and discarded the flow-through. After that we washed the QIAprep 2.0 Spin Column by adding 0.5 ml of Buffer PB and centrifuging for 1 minute, then discarding the flow-through. We then washed the QIAprep 2.0 Spin Column by adding 0.75 ml of Buffer PE and centrifuging for 1 minute and discarded the flow-through and centrifuged for an additional 1 minute to remove residual wash buffer. Then we placed the QIAprep 2.0 Spin Column in a clean 1.5 ml microcentrifuge tube. To elute DNA, we added 50 μl of Buffer EB (10 mM Tris·Cl, pH 8.5) and water to the centre of each QIAprep 2.0 Spin Column and let it stand for 1 minute.It was then centrifuged for 1 minute.

1.8.Quantification of DNA

Quantification of DNA was carried out using the Simplified QuantiFluor protocol. Samples were incubated at room temperature for 5 minutes before readings were taken. Standard readings were recorded as 204 ng/ul. For pSB1C30, we determined a concentration of 54 ng/ul and for pJUMP26-1A, a concentration of 16 ng/µL.

Digestion of Plasmids

Digestion of pSB1C30 and the pJUMP26-1A was carried out using EcoRI and PstI. The expected sizes of resulting fragments were 2029 bp for pSB1C30, and 2260 bp for pJUMP26-1A. The digestion mixture included 1 μl of DNA, 4 ul of 10x buffer, 1.5 μl of EcoRI, 1.5 μl of PstI, and 8 μl of water. Incubation occurred for 1 hour at 37°C for both plasmids.

Agarose Gel Electrophoresis

Gel electrophoresis was run at 10 V/cm, with an agarose gel prepared using 0.5 grams of agarose in 50 ml of TEAE 1X. Ethidium Bromide was added to the gel to visualise DNA. Electrophoresis was conducted using a Nahira gel box at 100 V. This step confirmed that the fragments were those desired as they corresponded to the expected sizes.

Fig.4- Digested DNA fragments on agarose gel electrophoresis

Gel Loading Dye (10x)

To facilitate loading DNA samples onto the agarose gel, a 10x gel loading dye was prepared. It consisted of 1 ml of DNA-loading Buffer, 500 μl of 50% glycerol (diluted in DEPC-treated water), 20 µl of 0.5 M EDTA (pH 8.0), and 480 μl of H2O. Additionally, 2.5 milligrams of bromophenol blue were dissolved in 100 millilitres of the solution, resulting in a concentration of 2.5 milligrams per millilitre of the dye solution.

Cloning Protocol

As a first-time iGEM team, we experienced problems with creating a cloning protocol that would actually be successful. Because of this delay, the DNA fragments didn’t arrive in time. We only had time to do the steps above, up to the point of the actual insertion of the genes into the plasmids (the last step we did was restriction digest of plasmid DNA), but we did thoroughly research the entire cloning strategy we proposed and we hope it will help future teams.

Cloning Strategy

Aims
Aim 1- Clone the transcription factor alkS under the control of a constitutive promoter
Aim 2 – Clone alkB1 under the control of alkS-inducible palkB promoter

Genes to clone
· alkS- Transcription factor involved in the expression of alkane-degrading genes. In the presence of alkanes, it binds palkB and inducing start of transcription.
· palkB – alkS-inducible promoter for alkB1
· alkB1 - Catalyzes the hydroxylation of n-alkanes and fatty acids in the presence of a NADH-rubredoxin reductase and rubredoxin.

A. Cloning constitutive promoter + alkS
· pJUMP26-1A (from the distribution kit plate 2 – well E17) as backbone (Kanamycin resistance)
· constitutive promoter J23119(order from IDT: prefix- J23119 -suffix)–suitable for bacterial and cyanobacterial systems
· alkS from IDT (prefix-RBS-alkS-suffix)

STEP 1: open pJUMP26-1A
1. Digest backbone pJUMP26-1A with EcoRI and PstI
2. Run gel electrophoresis of digestion reactions
3. Isolate part of interest by gel digestion
STEP 2: digest promoter J23119
4. Digest promoter J23119 with EcoRI and PstI
5. Clean the digestion reaction by PCR clean-up
STEP 3: digest alkS
6. Digest alkS with XbaI and PstI
7. Clean the digestion reaction by PCR clean-up
STEP 4: insert promoter J23119 in the open pJUMP26-1A plasmid
8. Ligate open pJUMP26-1A and J23119 using T4 ligase
9. Bacterial transformation
10. Miniprep
STEP 5: open pJUMP26-1A – J23119
11. Digest pJUMP26-1A – J23119 with SpeI and PstI
12. Clean the digestion reaction by PCR clean-up
STEP 6: insert alkS in the open pJUMP26-1A – J23119
13. Ligate alkS into pJUMP26-1A – J23119 using T4 ligase
14. Bacterial transformation
15. Miniprep

B. Cloning alkS-inducible promoter pAlkB + alkB1
· Plasmid pSB1C30 (from the distribution kit plate 1 – well O11) as backbone (Chloramphenicol resistance)
· alkS-inducible promoter pAlkB from IDT (prefix-pAlkB-suffix)
· alkB1 from IDT (prefix-RBS-alkB1-suffix)

Steps
1. Digest pSB1C30 with EcoRI and PstI
2. Digest pAlkB (from IDT; prefix-pAlkB-suffix) with EcoRI and PstI
3. Digest alkB1 (from IDT; prefix-RBS-alkB1-suffix) with XbaI and PstI
4. Run gel electrophoresis of digestion reactions
5. Isolate parts of interest by gel digestion
6. Ligate openpSB1C30 and pAlkB using T4 ligase
7. Bacterial transformation
8. Miniprep
9. Digest pSB1C30-pAlkB with SpeI and PstI
10. Clean the digestion reaction by PCR clean-up
11. Ligate open pSB1C30-pAlkB and AlkB1 with T4 ligase
12. Bacterial transformation
13. Miniprep

C. Cloning promoter J23119 + AlkS in plasmid with promoter pAlkB + AlkB1
· Plasmid pJUMP26-1A with promoter J23119 +AlkS (from step A)
· Plasmid pSB1C30 with pAlkB+alkB1 (from step B)

Steps
1. Digest plasmid pJUMP26-1A with promoter J23119 +AlkS with XbaI and PstI
2. Digest plasmid pSB1C30 with pAlkB+alkB1 with SpeI and PstI
3. Run gel electrophoresis of digestion reactions
4. Isolate parts of interest by gel digestion
5. Ligate open pSB1C30and pAlkB+alkB1 using T4 ligase
6. Bacterial transformation
7. Miniprep and sequencing(using standard primers VF2 and VR)

Standard protocols

1. Digestion
Add components to a clean tube in the order shown:
1. 1 µL DNA (concentration 1 µg/µL).
2. 2.5 µL 10x buffer CutSMART (or recommended buffer from NEB).
3. 1 µL each restriction enzyme.
4. 15 µL nuclease-free water.
5. Incubate the reaction at digestion temperature (usually 37 °C) for 1 hour.
6. Stop the digestion by heat inactivation at 65°C for 15 minutes (temperature may vary depending on which restriction enzymes were used).
7. At this point, samples may be stored at −20°C.
Alternatively, run DNA electrophoresis and proceed with gel extraction.

2. Ligation
Add components to a clean tube in the order shown:
1. Add 2 µL 10x reaction buffer for T4 DNA ligase.
2. Add 1 µL of T4 DNA ligase.
3. Add 60 ng DNA in total (mix of parts to ligate).
4. Add the necessary amount of nuclease-free water to give a final volume of 20 µL.
5. Incubate at room temperature (~22°C) for 30 min.
6. Heat-inactivate the T4 ligase by heating at 80°C for 20 min.
7. At this point, samples may be stored at −20°C.
Alternatively, run DNA electrophoresis and proceed with gel extraction.


3. DNA electrophoresis
Gel preparation - Casting a 150 ml gel
1. Set the DNA cast to the sealed position (depends on equipment)
2. For a 1% 150 mL agarose gel, weigh 1.5 g of agarose in a 500 mL conical flask. Add 150 mL 1x TAE/TBE buffer. If the gel tray is smaller, calculate the amount needed.
3. To dissolve the agarose in the buffer, swirl to mix and microwave for a few minutes taking care not to boil the solution out of the flask. Agarose has to be completely dissolve in the buffer and therefore, the solution should be transparent. In order to do this, make the solution boil for a few seconds, swirl and boil again. Be careful — the solution is very hot! Insulated gloves are too bulky to easily pull the flask out of the microwave, so a folded up paper towel is suggested.
4. Let the agarose solution cool down. Once the flask is touchable, add the DNA stain. The working concentration for Sybr®Safe is 1x (15 uL for 150mL gel).
5. Pour the gel solution into the gel tray. Remove any air bubbles with a pipette tip. Insert the comb into the gel tray at one end ~1 cm from the edge.
6. Let the gel solidify by cooling down to room temperature during ~30 min.

4. Run agarose gel
1. Release the gel tray from the casting stand. Place the gel tray into the buffer chamber and remove the comb carefully.
2. Add 1x TAE/TBE buffer until the gel is completely covered.
3. Take part of your DNA samples (~0.2 ug) and mix with loading dye. This can be done either in 1.5 mL tubes or, if the volumes are very small, on a piece of parafilm.
4. Load the size marker mixed in 1x loading dye (~6 uL final volume) into the first well. If you have many samples, you might load size markers in additional wells.
5. Load your samples into the other wells. Remember to write down which lanes have which samples.
6. Put the lid onto the buffer chamber and connect it to the power supply. Make sure to put it in the right direction so that your DNA runs towards the positive (red) electrode.
7. Run the gel at 100 V for 30-60 min. Neither of the two dyes should be run off the gel. If the electrophoresis runs correctly you will notice air bubbles coming from the negative (black) electrode.
8. Stop the run and bring your gel to a UV table to visualise your gel bands. Take a picture of your gel. If sufficient separation of the bands was not achieved, put the gel back into the buffer chamber and run it for longer.

5. Gel digestion
1. Take your gel to the UV table to visualise your gel bands.
2. Take a scaple and cut out the bands of interest. Remove as much agarose as possible.
3. Put each band in an eppendorf tube.
4. Proceed with gel digestion protocol following kit instructions.
5. Elute with 20 uL elution buffer or nuclease-free water.
6. Measure concentration and purity of eluted DNA.

References

[1] Josefine Liljeruhm, Erik Gullberg, Anthony C. Forster. Synthetic biology. A lab manual