Experiments & Protocols

In Vivo Assembly (IVA) Cloning

Overview. This protocol assembles DNA constructs into the pEM106.2 backbone using In Vivo Assembly cloning. The method uses the homologous recombination machinery of E. coli to join linear backbone and insert fragments that share overlapping ends, replacing the YebF-hIL10 cassette with our FAAH constructs. No restriction enzymes or ligase are needed.

Because backbone and insert amplify in separate reactions, our six-construct panel ran as twelve PCR reactions. Both numbers appear in our records and they describe different things.

Materials

• Template DNA: pEM106.2 backbone, and inserts (FAAH, GFP, OmpA, OmpX)
• Primers: custom IVA primers with 15–20 bp overlaps
• Enzymes: Phusion High-Fidelity DNA Polymerase, DpnI
• Buffers: 5× Phusion HF Buffer, 10 mM dNTPs
• Competent cells: E. coli Stable or DH5α
• Selection: LB agar with chloramphenicol at 25 µg/mL

PCR reaction mix (25 µL)

Step by step

1. Set up separate reactions for the backbone and the insert using the matched primer pairs.
2. Run the thermal program: 30 s at 95 °C; 18 cycles of 10 s at 95 °C, 30 s at 60 °C, 4 min at 72 °C; 5 min final extension at 72 °C; hold at 4 °C.
3. Add 1 µL DpnI to each reaction and incubate 30 min at 37 °C to digest methylated parental template.
4. Run 5–10 µL on a 0.5–1% agarose gel to confirm the fragment size.
5. Mix equal molar amounts of backbone and insert product and transform directly into competent cells by heat shock.
6. Plate on LB agar with 25 µg/mL chloramphenicol and incubate overnight at 37 °C.
7. Pick colonies for miniprep, screen by restriction digest, and confirm by Sanger sequencing.

Gradient PCR for Troubleshooting

Overview. When a reaction gives no product, multiple bands, or a smear, we re-run it across an annealing temperature gradient. This is the single technique that rescued the most reactions for us.

Step by step

1. Split the reaction across 7–8 tubes.
2. Set an annealing gradient. Our standing fix, set on 18 June, was 55–65 °C. In practice we ran 55–67 °C for insert amplification and 61–68 °C for whole-plasmid amplification, and the wider insert range is what we would use again.
3. Raise the cycle number from 18 to 20 if the product is faint. The 25 June gradient was run at 17 cycles; 20 cycles on 29 June gave the usable product.
4. Run all tubes side by side on one 0.5% gel with a 1 kb ladder so the temperatures can be compared directly.
5. Carry forward the temperature that gives a single band at the expected size.

Agarose Gel Electrophoresis

Overview. Standard size separation of PCR products and restriction digests.

Step by step

1. Prepare a 0.5–1% agarose gel in 1× TAE. Use 0.5% for fragments above 3 kb.
2. Add SYBR Safe to the molten gel, or use ethidium bromide where a more sensitive stain is needed.
3. Pour into the casting tray and leave 30 min to set.
4. Cover with 1× TAE buffer.
5. Mix DNA with loading dye and load 10 µL per well alongside a 100 bp or 1 kb ladder.
6. Run at 160 V for 30 min. Reduce voltage for larger fragments needing better resolution.
7. Image under UV.

Gel Extraction

Overview. Recovery of a single correct band when a PCR gives more than one product.

Step by step

1. Run the product on a 0.5% gel at low voltage to maximize separation between close bands.
2. Image, then excise the target band with a clean blade under long-wave UV, minimizing exposure time.
3. Purify with a commercial gel extraction kit following the manufacturer’s instructions.
4. Quantify on a Nanodrop before proceeding.

Site-Directed Deletion by In Vitro Ligation

Overview. This protocol removes an unintended extra ‘A’ nucleotide at the junction of the YebF-ΔTm-FAAH construct, clones #19 and #21. We first tried to do this by amplifying the whole plasmid and letting E. coli circularize the blunt product. That failed three times across eight weeks, from the sequencing that found the extra base on 20 July to the sequencing that confirmed it gone on 15 September. The protocol below, using in vitro phosphorylation and ligation, worked on the first attempt.

Materials

• Template: YebF-ΔTm-FAAH plasmid carrying the extra ‘A’
• Primers: 19_delA_Fw (CGAGCTGGCCACCCTCTCGTC) and 19_delA_Rv (TTACGCCGCTGATATTCCGCCATTCC)
• Enzymes: Phusion polymerase, DpnI, T4 Polynucleotide Kinase, T4 DNA Ligase
• Gel extraction kit
• E. coli Stable competent cells

Step by step

1. Amplify the entire plasmid with the deletion primers, which anneal either side of the unwanted base so it is excluded from the product. Optimize annealing across 61–68 °C.
2. Treat the product with DpnI overnight at 37 °C. An overnight digest matters here: a short digest leaves enough parental plasmid to dominate the transformation and return only parent sequence.
3. Run on a gel and excise the ~3 kb band. Purify with a gel extraction kit.
4. Phosphorylate the 5’ ends with T4 PNK. PCR products carry no 5’ phosphate, so without this step ligation cannot proceed.
5. Circularize with T4 DNA ligase.
6. Transform into E. coli Stable cells.
7. Pick 5 colonies, miniprep, and sequence across the junction.

Controls

• Negative control: PCR with no template, to detect contamination.
• Parent plasmid sequenced alongside, so an unchanged clone is recognisable immediately.

Restriction Digest Screening

Overview. A fast first-pass check on miniprep DNA before paying for sequencing.

Step by step

1. Map restriction sites in the insert and the backbone, and choose an enzyme pair that gives a clearly different band pattern for correct and empty vector.
2. Prepare a mastermix per reaction: 6.5 µL buffer, 6 µL enzyme, 6.5 µL nuclease-free water.
3. Add 2 µL miniprep DNA per reaction.
4. Incubate at the enzyme’s recommended temperature.
5. Run on a 1% agarose gel against an uncut control.

Heat Shock Transformation

Overview. Introduction of assembled plasmid, or of linear IVA fragments, into competent E. coli.

Step by step

1. Thaw competent cells on ice.
2. Add the DNA, or the mixed backbone and insert products, and incubate on ice for 30 min.
3. Heat shock at 42 °C for 45 s, then return to ice for 2 min.
4. Add recovery medium and shake at 37 °C for 1 hour.
5. Plate on LB agar with 25 µg/mL chloramphenicol.
6. Incubate overnight at 37 °C.

Overnight Culture and Selection

Step by step

1. Label one tube per colony and add 4–5 mL LB broth with 25 µg/mL chloramphenicol.
2. Pick a single colony with a sterile tip and inoculate.
3. Incubate at 37 °C with shaking at 120 rpm overnight.
4. Check for turbidity before proceeding to miniprep.

Miniprep

Step by step

1. Pellet the overnight culture by centrifugation and discard the supernatant.
2. Resuspend, lyse, and neutralize following the kit protocol.
3. Clear the lysate and load onto the silica spin column.
4. Wash, then elute into a clean labelled Eppendorf tube.
5. Quantify on a Nanodrop.

Protein Expression in E. coli BL21(DE3)

Overview. Induction of 6xHis-tagged FAAH variants for expression analysis.

Step by step

1. Inoculate a single colony into 5 mL LB with chloramphenicol. Grow overnight at 37 °C, 120 rpm.
2. Dilute 1 mL of overnight culture into 100 mL fresh LB with chloramphenicol.
3. Grow at 37 °C to OD₆₀₀ of 0.6–0.8.
4. Induce with doxycycline at 1 µg/mL final concentration.
5. Incubate 3 hours at 37 °C.
6. Harvest the cells by centrifugation. Store the pellet at −20 °C.
7. Retain an uninduced sample taken before step 4. Without it the induced lanes cannot be interpreted.

SDS-PAGE

Overview. Size separation of total cell protein to confirm induced expression.

Step by step

1. Resuspend the cell pellet and lyse.
2. Separate into pellet and supernatant fractions by centrifugation.
3. Mix each fraction with loading buffer and heat to denature.
4. Load uninduced, induced pellet, and induced supernatant for each construct side by side, with a molecular weight ladder.
5. Run until the dye front reaches the bottom of the gel.
6. Stain, destain, and image.

Expected sizes

Western Blot Against the 6xHis Tag

Overview. Confirms that the induced band seen by SDS-PAGE is the tagged protein. In progress at the time of writing.

Step by step

1. Run SDS-PAGE as above.
2. Transfer to a membrane.
3. Block, then probe with an anti-6xHis primary antibody.
4. Wash, apply the labelled secondary antibody, and wash again.
5. Develop and image.

TAMRA-FP Active Site Labelling

Overview. Activity-based probe labelling that reports whether the FAAH serine hydrolase active site is intact and accessible. Planned.

Step by step

1. Incubate cell lysate with the TAMRA-fluorophosphonate probe, which binds covalently to active serine hydrolase sites.
2. Quench and separate by SDS-PAGE.
3. Image the gel for TAMRA fluorescence before staining for total protein.

Controls

• Catalytic mutant FAAH, which should give no labelled band. This is the control that makes the result interpretable.
• Heat-inactivated lysate as a second negative.

Full detail is on our Safety and Security page. Two reagents in the protocols above need direct mention here.

• Ethidium bromide is a mutagen and an irritant. We use SYBR Safe as the default stain and reserve ethidium bromide for cases where sensitivity demands it. When it is used: gloves at all times, a designated staining tray and bench area, no shared equipment contact, and disposal of gels and gloves as hazardous waste.
• Ultraviolet light damages skin and eyes. Band excision is done with the shortest possible exposure, with the face shield down and arms covered.

All cultures, plates, tips, and liquid waste go into autoclave waste. No engineered strain leaves the laboratory. Our chassis organisms, E. coli Nissle 1917, BL21(DE3) and Stable, are all non-pathogenic laboratory or probiotic strains handled under BSL-1 conditions.