In this protocol,you can find how to perform Gibson cloning.

Materials:

• Gibson master mix.
• Backbone fragment.
• Insert fragment.

Protocol:

1. For Gibson plasmid assembly prepare the following reaction mixture in a 100 µL PCR tube according to figure 1.


Figure 1. Overview of the composition of the Gibson cloning reaction mixture.

2. Mix gently and incubate at 50°C for 45-60 minutes.

If the Gibson master mix isn’t available, make new according to figure 2.

1. Prepare 5x isothermal reaction buffer. Recipe for 4 mL (don’t combine materials immediately. Read the steps):
2. Mix dNTPs, NAD+, Tris-HCl, MgCl2 and DTT.
3. Slowly add PEG-8000 to the mixture and mix well until completely dissolved. Add H2O to a final volume of 4 mL.
4. Prepare aliquots of the 5x isothermal buffer as required. Store at -20C.
5. Prepare 1,33x Gibson Assembly Mastermix (figure 3). Recipe for 25 x 15 uL aliquots (don’t combine materials immediately. Read the steps):
6. Work on ice. Mix H2O and 5x buffer, then add enzymes.
7. Prepare 25 x 15 uL aliquots in PCR tubes. Store at -20C.

Note: The aliquots are concentrated in 1,33x. Add DNA in a volume of 5 uL to a final volume/concentration of 20 uL/1x.

Figure 2. Overview of the composition of 5x isothermal reaction buffer.

Figure 3. Overview of the composition of the Gibson master mix.

In this protocol, you can find how to perform Golden Gate cloning.

Materials:

• Golden Gate master mix.
• Backbone fragment.
• Insert fragment.
• T4 ligase buffer.
• T4 ligase
• MilliQ

Protocols:

1. For plasmid assembly prepare a reaction mixture in a 100 µL PCR tube according to figure 4.


Figure 4. Overview composition of Golden Gate reaction mixture.

2. Mix gently by pipetting up and down.
3. Place the tubes in a PCR machine and follow the protocol from figure 5.


Figure 5. PCR protocol Golden Gate reaction.

In this protocol,you can find how to perform a polymerase chain reaction (PCR).

Materials:

• Forward and Reverse primer for your DNA plasmid.
• Phusion buffer (5x).
• dNTPs.
• MilliQ.
• Phusion DNA polymerase.

Protocol:

1. Prepare PCR mixture based on figure 6.
2. The PCR mix is mixed by gently pipetting up and down, followed by insertion into a PCR machine with the program depicted in figure 7.
3. Primer annealing temperature is determined using Snapgene for each primer. Then the found Tms are chosen minus 3-5°C. What amount is subtracted depends on the difference in Tm between the Fw and Rev primers. E.g. a primer of Tm= 61°C and one of Tm= 59°C will be run at an annealing temperature of 56°C.
4. Primer extension depends on the desired length of the fragment, with 30 seconds per kb of product length.

Figure 6. PCR mixture.

Figure 7. Touchdown PCR program.

In this protocol, you can find how to perform a colony polymerase chain reaction (PCR).

Materials:

• Forward & Reverse primer for your DNA plasmid.
• Phire Hot Start II PCR Master Mix.
• MilliQ.

Protocol:

1. Resuspend a colony in 20 μL sterile water.
2. Prepare the PCR mixture by adding together 10 uL of Phire hot start 2x, 1uL forward primer, 1 uL reverse primer, and 7 uL dH2O.
3. To the PCR mixture add 1 uL of the resuspended colony mixture.
4. Primer annealing temperature is determined using Snapgene for each primer. Then the found Tms are chosen minus 3-5°C. What amount is subtracted depends on the difference in Tm between the forward and reverse primers. E.g. a primer of Tm= 61°C and one of Tm= 59°C will be run at an annealing temperature of 56°C.
5. Primer extension depends on the desired length of the fragment, with 15 seconds per kb of product length.

Figure 8. Touchdown PCR protocol for colony PCR.

In this protocol, you can find how to prepare samples for sequencing.

Materials:

• GeneJET Plasmid Midiprep Kit.
• 1,5 ml tubes for each sample respectively.
• Overnight culture of bacteria containing DNA plasmid.

Protocol:

1. Perform Plasmid Midiprep Kit.
2. Perform Nanodrop on obtained samples to determine the DNA concentration.
3. Add 5 µL of plasmid solution to a 1,5 mL tube.
4. Add 5 µL of 10 uM forward & reverse primer, for the plasmid respectively.
5. Label tubes and send them for sequencing.

In this protocol, you can find how to perform an Agarose gel electrophoresis, to separate DNA samples based on their size.

Materials:

• Agarose.
• SERVA DNA Stain G (working dilution is 1:20,000 to 1:50,000).
• TAE buffer.

Protocol:

1. Weigh x% m/v of agarose gel and place this in a flask.
1. When 20 mL of 1% agarose is made, 0.2 grams of agarose is weighed.
2. When 20 mL of 3% agarose is made, 0.6 grams is weighed, etc.
2. Add the TAE buffer and dissolve using a microwave.
1. When microwaving be sure to place a piece of paper (not the writing type, the toilet- or kitchen variant) underneath the flask in case of spilling.
2. Keep an eye on it and don’t let it boil too much.
3. Safety warning: From here on out, equip 2 layers of (blue) gloves and be sure not to touch anything which has come into contact with Ethidium Bromide (EtBr) with bare hands. It’s carcinogenic. Double gloves allow you to remove a layer if you touch any gel/EtBr directly without risking contact.

Note: we used SERVA DNA Stain G, which can also be used in place of EtBr, which is less risky but still caution is advised.

1. Also means that you can only keep anything contaminated with EtBr in the designated area. This includes the gel, the gloves, the pipettes. A special trash-can will be placed nearby for waste.
1. One exception is the gel when you go for viewing, as this is unavoidable. But even then, don’t touch any part of the machine except the glass gel-plate with contaminated gloves. Also clean everything afterwards with paper and water.
2. When you get into direct contact with EtBr, immediately wash thoroughly with water and soap and you should be fine.
3. Gels are poured into a mold followed by the immediate addition of 4 µL of ethidium bromide, which is mixed using the pipet tip as a stirrer.
4. Don’t forget to add a comb.
4. Wait for solidification, which may take around 20 minutes.
5. Place the gel (with the see-through support but without the red barriers) into the Electrophoresis Reservoir.
6. Gently remove the comb and add 1x TAE to the reservoir. The TAE can be re-used a few times, but try to replace it once every day or two to keep the gel free from contamination.
7. Carefully add the sample + dye to the wells.
1. Dye is often 6x concentrated, so for 5 µL samples you’d need to add 1 µL dye, for 20 µL samples you’d need 4 µL dye, etc. Dye can be made by ourselves, or bought, varying the concentrations.
2. The volume per well depends on the comb used, but can be anywhere from max. 15 µL to 50 µL. You could do some testing to find out the maximum.
3. Tip: When you have the sample + dye in your pipet tip, the moment it touches the TAE some of the sample may come out. So penetrate the surface at the bottom deeper part of the reservoir, then move to the wells. This way you don’t accidentally spill any sample in the wrong well. Then after gently releasing the sample into the well, move up a bit and suck in 1 µL of the TAE so you don’t drag up the sample while moving out of the liquid again.
8. Close the machine and be sure to connect the wires correctly.
1. The positive electrode pulls on the negatively charged DNA, so have the positive electrode at the bottom, and the negative electrode on the well-side.
9. Run the electrophoresis at 80 V until the dyes have traveled through 80% of the gel, which will take 40-60 minutes. Higher voltages are possible to speed up the process, but this may melt the gel if you do it too much.
10. After electrophoresis, turn off the machine, remove the lid, and carefully lift the gel out of the TAE.
1. The gel is not stuck on the support, so it may float. Be sure you prevent that.
11. Let the liquid seep out and remove the gel from the support with a double-gloved hand.
12. Carefully remove one layer of gloves from one hand, so the newly revealed glove is uncontaminated, and use this hand to open the UV imager machine.
1. Then image the gel, followed by removal of the gel and cleaning of all materials.

In these two protocols, you can find two ways to make competent Escherichia coli DH5𝛂 or E.coli BL21 cell aliquots.

Materials 1:

• LB media.
• Ice-cold 0.1 M CaCl2.
• Ice-cold 0.1 M CaCl2 + 10% glycerol.
• Sterile tubes (1,5 ml).

Protocol 1:

1. Inoculate the E.coli DH5a in 2 mL LB medium for overnight growth at 37℃ and at 220 RPM.
2. After overnight growth, inoculate fresh medium in a sterile erlenmeyer flask with a volume ratio of 1/100. Usually this means 20 mL LB with 0.2 mL of the overnight culture, but it can be scaled up, which is recommended when an -80℃ freezer is available.
3. Grow at 37℃, 220 RPM until an OD of 0.4-0.6. Measure the OD with a spectrophotometer.
1. When the desired OD is approaching, pre-cool a centrifuge.
4. Transfer the culture to a sterile 50 mL tube and spin down in a pre-cooled centrifuge, at 5℃ for 10 minutes at 5000 RPM.
5. Discard the supernatant and add 0.5*[Culture Volume] of ice-cold 0.1 M CaCl2. Resuspend carefully and incubate on ice for at least 30 minutes.
6. Spin down the cells in a pre-cooled centrifuge at 4℃ , for 10 minutes at 5000 RPM. Discard the supernatant carefully.
7. Resuspend the cells in 0.1*[Culture Volume] ice-cold 0.1 M CaCl2 + 10% glycerol. Aliquot in 100 µL fractions in sterile 1.5 mL tubes stored at -80 (or -20)℃

Materials 2:

• TYM media:
• 20 g/L bacto-tryptone.
• 5 g/L Yeast extract.
• 0.1 M NaCl.
• 10 mM MgSO4.
• TfB I:
• 30 mM KAc.
• 50 mM MnCl2 (sterilize through filter, add later).
• 0.1 M KCl.
• 10 mM CaCl2.
• 15% glycerol.
• TfB II:
• 10 mM Na-MOPS, pH 7.0.
• 75 mM CaCl2.
• 10 mM KCl.
• 15% glycerol.

Protocol 2:

1. Inoculate a colony in TYM medium.
2. Grow cells to midlog phase (OD600 = 0,2-0,6).
3. Transfer cells to 100ml of TYM.
4. Grow cells to OD600 = 0,5-0,9.
5. Add TYM medium to 500ml.
6. Grow cells to OD600 = 0,6.
7. Cool cell culture on ice (keep cells cold from now on).
8. Centrifuge 15min, GSA rotor, 4200rpm.
9. Resuspend pellet in 100ml of cold TfB I (on ice).
10. Centrifuge 10min, GSA rotor, 4200rpm.
11. Resuspend pellet in 20ml of cold TfB II (on ice).
12. Aliquot in 300µL (use 150µL per transformation).
13. Freeze in liquid nitrogen.
14. Store at -80°C.

In this protocol, you can find how to make competent Lactobacillus reuteri DSM20016 cells for immediate use.

Materials:

• MRS media.
• MRS broth containing 2% glycine and 0.5 M sucrose.
• Cold deionized water.
• 50 mM EDTA (pH 8.0)
• 0.3 M sucrose

Protocol:

1. Grow L. reuteri overnight at 37˚C in MRS broth
2. Inoculate 15 mL MRS broth containing 2% glycine and 0.5 M sucrose with the overnight L. reuteri culture. The glycine acts as a cell wall weakening agent.
3. Grow the cells at 37˚C to an OD600 of 0.2 (~3 hours).
4. Spin down the cells at 6000 g for 10 minutes at 4 ˚C
5. Wash the cells twice with 5 mL cold distilled water
6. Incubate the cells in 50 mM EDTA (pH 8.0) for 10 min
7. Wash the cells with 0.3 M sucrose
8. Resuspend the cells in 100 uL 0.3 M sucrose as electroporation buffer

In this protocol, you can find how to transform competent E.coli cells. (how to insert plasmid DNA into your bacteria).

Materials:

• Ice
• Prewarm heat block or water bath at 42ºC
• Selective plates with the corresponding antibiotics
• Competent cells
• Plasmid DNA

Protocol:

1. Defrost 150 µl aliquot of E. coli competent cells per transformation on ice.
2. Add 1 μl plasmid DNA (10 pg/μl), incubate 15 min on ice.
3. Heat shock the cells for 1 min at 42°C.
4. Immediately place the cells on ice for 2 min.
5. Add 1mL LB and incubate at 37°C for 1 hour (200rpm).
6. Plate out on selective plates.
7. Incubate plates overnight at 37ºC.

In this protocol, you can find how to transform competent L.reuteri cells. (how to insert plasmid DNA into your bacteria).

Materials:

• Recovery medium (sterile, kept at 4°C):
• M17 (amount dependent on volume).
• 0.5M sucrose.
• 0.5% glucose.
• 20 mM MgCl2 (prepared as stock and sterilized separately).
• 2 mM CaCl2 (prepared as stock and sterilized separately).
• GM17-agar (sterile and poured in standard petri-dishes with appropriate antibiotics):
• M17 (amount dependent on volume).
• 0.5% glucose.
• 1.5% agar.

Protocol:

1. Thaw competent L. lactis cells on ice.
2. Add a maximum of 10 µL of the DNA to be transformed to the competent cells. Immediately continue to the next step.
3. Transfer the cell/DNA mixture to a sterile electroporation cuvette.
4. Place the cuvette in the electroporation machine and run for approximately 4-5 seconds at the following settings: 2.25 kV, 25 µF and 200 Ω.
5. Add 1 mL of recovery medium to the electroporation cuvettes and carefully mix, then transfer the suspension to 2 mL tubes.
6. Incubate for 2 hours at 30°C.
7. Plate 100 µL of the suspension per plate. It is also possible to then pellet the cultures and resuspend in lower volumes to concentrate the cells, so more colonies could form. This is at risk of covering the plates too much, making differentiation between colonies impossible.

In this protocol, you can find how to make LB media.

Materials:

• LB (lennox) powder.
• Deionized water.

Dissolve 20 grams of LB-medium powder in 1 liter of deionized water. Autoclave bottle and close lid when cooled down.

Composition:

• NaCl, 5 g/L.
• Tryptone, 10 g/L.
• Yeast extract, 5 g/L.

In this protocol, you can find how to make MRS media.

Materials:

• MRS (Millipore) powder.
• Deionized water.

Dissolve 51 grams of MRS-medium powder in 1 liter of deionized water. Before autoclaving, preheat the autoclave to limit the glucose from caramelizing. Autoclave bottle and close lid when cooled down.

Composition:

• Peptone 10,0 g/L.
• Meat extract 8,0 g/L.
• Yeast extract 4,0 g/L.
• D(+)-Glucose 20,0 g/L
• Dipotassium hydrogen phosphate 2,0 g/L.
• Sodium acetate trihydrate 5,0 g/L.
• Triammonium citrate 2,0 g/L.
• Magnesium sulfate heptahydrate 0,2 g/L.
• Manganous sulfate tetrahydrate 0,05 g/L.
• Final pH 6.2 +/- 0.2 at 25°C.

In this protocol, you can find how to make M9 media.

Composition for 1 liter:

• 200 mL 5x M9 salts (Sigma Aldrich), autoclaved.
• 2 mL 1M MgSO4 (Thermofisher), autoclaved.
• 100 uL 1M CaCl2 (Thermofisher), autoclaved.
• 780 mL dH2O, autoclaved.
• 20 mL 20% Glucose, filter sterilized (add after autoclaving).

For the creation of Agar plates with a certain media, all you have to do is add 15 grams of agar to 1 Liter of media respectively before autoclaving. Keep the media at 60 degrees Celsius after autoclaving to prevent the media from stolling. If you want to add selective antibiotic markers, let the media cool down until you can hold the bottle comfortably for over a minute in a gloveless hand and add the antibiotics before pouring the plates. Use about 15-20 ml per plate.

In this protocol, you can find how to perform an ELISA experiment.

Materials:

• 96 well plate (maxisorp plates from nunc).
• Desired hemagglutinin proteins.
• PBS.
• PBS with 3% (w/v) skimmed milk powder.
• PBS with 0,05% (v/v) Tween-20.
• Anti c-myc 9E10-HRP.
• OPD.
• 2 M sulphuric acid.

Protocol:

1. Coat the plates with 0.1 mL hemagglutinin at 1 µg/mL overnight in PBS at 4 ˚C.
2. Wash the wells 3 times with 300 µL PBS.
3. Block the wells with 200 µL PBS 3% (w/v) dried milk protein for 1 hour at room temperature.
4. Wash the wells 3 times with 300 µL PBS containing 0.05% (v/v) Tween-20.
5. Add crude extracts (25% v/v) or purified nanobody (100 µL) to the plate and incubate for 1 hour at room temperature.
6. Wash 5 times with 300 µL PBS containing 0.05% (v/v) Tween-20.
7. To detect antibody binding 100 µL of anti c-myc 9E10-HRP (1/1000 dilution) (Roche) in 2%.
8. (w/v) milk powder in PBS was added. Plates were incubated for 1 hour at room temperature.
9. Wash the wells 3 times with 300 µL PBS containing 0.05% (v/v) Tween-20.
10. Add 100 uL chemiluminescent reaction (OPS).
11. Stop the reaction by adding 50 uL 2 M sulphuric acid.
12. Read the plate at 450 nm.

In this protocol, you can find how to prepare and perform a sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS PAGE) protein separation.

Materials:

• 30% Acrylamide/Bis Solution 37.5:1 (2.6%C).
• TemeDel.
• 10% Ammonium persulfate (APS).
• 25 mM Tris, 192 mM glycine and 0.1% SDS, with pH adjusted to 8.7 (running buffer).
• 25 mM Tris, 192 mM glycine and 0.1% SDS, with pH adjusted to 6.8 (stacking buffer).
• Iso-propanol.

Protocol:

PART I: pouring the gel.

1. Prepare running gel according to figure 9.
2. Pour the gel with a pipet between the glasses for the SDS Page, leave enough room for the stacking gel on top and the comb (which should hover about half a centimeter above the running gel.
3. To remove bubbles and create a flat top of the gel pipet iso-propanol on top of the gel. (discard after the gel has solidified)
4. Prepare stacking gel according to figure 9
5. Pour the gel with a pipet to the top and place the comb for the slots.
6. After the stacking gel has solidified, remove the comb straight and smooth in one go. Then place the gel in the gel electrode assembly with the smaller glass facing inwards.

Part II: running the gel.

7. Fill the inside of the holder to the top with a running buffer and ensure the electrode at the bottom is submerged.
8. Load the samples with the loading dye.
9. Run the gel at 100 volt for about an hour.
10. Stain the gel afterwards with coomassie blue staining for about 20-30 min (this process can be sped up by microwaving the gel shortly)
11. Destain the gel overnight while submerged in deionized water and shaking.

Figure 9. Showing composition of running and stacking gels according to desired percentage of Acrylamide.

In this protocol, you can find how to perform a Western blot used to visualize certain proteins.

Materials:

• Whatmann papers cut to squares the size of the gel (6 for 1 gel).
• 2 sponges (for 1 gel).
• 100 mL methanol.
• Ice pack filled with frozen blotting buffer.
• PVDF or nitrocellulose membrane for western blot (always wear gloves when handling!).
• 1X PBS:
• 4 mM KH2PO4.
• 14 mM Na2HPO4.
• 115 mM NaCl.
• 1X PBS-T.
• Add 0.05% Tween20 to PBS
• Fresh 1X blot buffer (2L).
• 14.4 g Glycine.
• 3g Tris.
• Add dH20 up to 850 mL.
• Add 150 mL methanol.
• Blocking buffer.
• 0.2% I-block in PBS-T. Warm up PBS and dissolve I-block in there, then add Tween. Tween cannot be heated.
• SuperSignal West Pico Plus Chemiluminescent Substrate.
• Fuji imager.

Protocol:

PART I: Protein transfer to membrane.

1. Incubate a SDS-PAGE gel for 15 min in blot buffer, as well as 3 stacks of Whatmann papers.
2. Incubate the membrane for 15 min in methanol and dip it in blot buffer. Make sure the membrane does not dry out during assembly of the sandwich!
3. Assemble the sandwich in the following order in the blotting casette:
1. Black side (-).
2. Sponge
3. 3x Whatmann paper.
4. SDS-PAGE gel.
5. Membrane.
6. 3x Whatmann paper.
7. Sponge
8. Red/White side (+).
4. Remove air bubbles by rolling over the sandwich (i.e. with an empty falcon tube).
5. Close the sandwich and transfer to a running tank, black on the black side. Add blot buffer and an ice pack (filled with frozen blotting buffer).
6. Run for 1.5h at 100V.
7. Carefully take out your membrane and put it in a 50ml falcon tube using tweezers (make sure it does not dry out!). The protein side of the membrane should be facing to the inside of the tube.
8. Swiftly fill the falcon tube with blocking buffer.
9. Block your membrane overnight in the cold room, or for 1h at RT. Remember that the falcon tube should always be shaking/turning.

PART II: Visualization

1. Discard the blocking buffer buffer and shortly rinse the membrane with PBS-T in a squared petri dish.
2. Add 1:10.000 antibody (anti-his-HRP) diluted in 0.1% I-block in PBS-T (15 mL for one blot in a square petri dish) and incubate for 1h shaking at RT.
3. Collect the antibody and store it in -20C, it can be used several times.
4. Wash the membrane 3x 10 min shaking at RT with PBS-T.
5. Prepare 1mL of SuperSignal West Pico Plus Chemiluminescent Substrate (500ul of each component).
6. Take the membrane out of the PBS-T, drain of excess liquid and transfer the membrane into a plastic sheet. Add the 1mL of substrate on top of the membrane.
7. Image the blot in the Fuji imager by chemiluminescence/increments on a black background and -without moving the blot- also take an image of the protein ladder in the EPI channel.

In this protocol, you can find how to perform Sonification used to break cells apart.

Materials:

• Sonicator.
• Lysis buffer.
• Ice.

This protocol begins when a culture of cells is available.

1. Centrifuge cells to pellet them (5000 RPM for 5-10 minutes for moderate pelleting). Discard the supernatant (or in the case of a protein secretion experiment, save the supernatant for further testing).
2. Resuspend the cells in a lysis buffer.
1. Usually contains PMSF (phenylmethylsulfonyl fluoride) which prevents protein degradation.
3. Chill the cell solution in ice, and keep the sample in ice throughout the process.
1. Sonication generates heat which might damage proteins.
4. Put the sonication probe inside the tube with the chilled resuspended cells.
1. Make sure the bottom of the probe is fully submerged, but don’t let it touch the bottom of the tube.
2. Keep an eye on the sonication, as foaming indicates protein denaturing.
5. Determine the parameters of the sonication.
1. Frequency: Often set between 20-50 kHz, depending on the lysis difficulty and protein sensitivity.
2. Sonication duration varies depending on optimization, but is often from 10-30 seconds.
3. Duration of intervals varies, but is important for cooling down between steps. Can be from 10-60 seconds.
4. One protocol suggests: Sonication until 400J using 50% amplitude and pulses of 10s: 10s (so 10 second duration with 10 second intervals).

In this protocol, you can find how to perform a His-tag purification.

Materials:

• Ni-NTA sepharose resin (beads).
• 1x phosphate buffered saline (PBS).
• 2M imidazole.

Protocol:

1. Add 1 mL Ni-NTA sepharose resin to the column (0.5 mL column volume).
2. Wash beads with 10 CV deionized water.
3. Wash beads with 10 CV 1x PBS + 10 mM imidazole.
4. Add the washed beads to the cell free extract containing the protein of interest. Incubate the cell free extract with the Ni-NTA beads for 1 hour at 4 °C under constant nutation.
5. Add protein & Ni-NTA beads mixture to the column to establish the column. Collect a sample for the flowthrough.
6. Wash the column with 20CV PBS + 50mM of imidazole. Collect a sample from the washing step.
7. Elute the column with PBS + 500mM imidazole. Collect 15 fractions of 150 µl each.

In this protocol, you can find how to perform a Bradford assay.

Materials:

• Coomassie brilliant blue G-250 dye.
• 85% phosphoric acid.
• 95% ethanol.
• Sterile MilliQ.
• BSA.

Protocol:

Below is the GoldBio protein quantification assay.

Bradford reagent preparation:

1. Weigh 100 mg coomassie brilliant blue G-250 dye.
2. Add 50 mL 95% ethanol.
3. Slowly and carefully add 100 mL 85% phosphoric acid.
4. Mix until the blue dye is completely dissolved.
5. Add the dye/ethanol/phosphoric acid solution to 850 mL sterile MQ H2O.
6. Filter any precipitates.
7. Store Bradford Reagent at 4℃ for months.

Measuring Protein Standard:

1. Warm up the spectrophotometer.
2. Pipet 6 different volumes of 0.5 mg/mL BSA into separate cuvettes.
1. E.g.: 0 µL (0 µg), 10 µL (5 µg), 20 µL (10 µg), 30 µL (15 µg), 40 µL (20 µg), 50 µL (25 µg).
2. Other protein standards can be substituted for BSA.
3. The cuvette with no protein standard in it serves as a blank.
3. Add 1.5 mL of Bradford Reagent to each cuvette.
4. Cover the cuvettes with plastic paraffin film and mix by gently inverting.
5. Let the cuvettes incubate at room temperature for 10 minutes.
6. Measure the absorbance of each cuvette at 595 nm.

Measuring the sample of unknown quantity:

1. Pipet between 10 to 50 µL of the protein that is to be quantified into a cuvette; a dilution may be necessary.
1. The goal is to get an absorbance reading that is in the middle of the standard curve.
2. Add 1.5 mL of Bradford Reagent to the cuvette.
3. Cover the cuvette with plastic paraffin film and mix gently by inverting.
4. Let the cuvettes incubate at room temperature for 10 minutes.
5. Measure the absorbance of each cuvette at 595 nm.
1. If the absorbance doesn’t fall within the range of the standard curve, change the dilution of the sample and measure the absorbance again.

Standard Curve Generation:

1. Make a scatter plot for the standard curve values.
2. Plot the micrograms (µg) of standard protein that was assay on the x-axis. Plot the Abs595 of the measurements on the y-axis.
3. Fit a linear trendline to the graph.
4. Show the linear regression equation.
5. Calculate the unknown amount of protein by using the linear regression equation, as shown in the below picture (figure 10):


Figure 10. Formula used to calculate the unknown amount of protein: the linear regression equation.

This is the DNA extraction protocol we initially used for gathering the DNA from the lungs. It is based on the homogenization strategy of a tissue sample described in [1]. Keep in mind that for analysis of microbiomes without many microbes living in it (e.g. lungs), your microbial DNA concentration might end up too low for further downstream analysis. For this we recommend our second DNA extraction protocol from a tissue sample (not suitable for studying percentages in microbial compositions, only for the presence of certain microbes).

Materials:

• Chicken lungs (acquired as a by-product from the meat industry)
• Liquid nitrogen
• Mortar and pestle
• Biosafety cabinet
• DNeasy PowerSoil Pro Kit

Protocol: an overview of the steps taken can be found in figure 10.

• Lungs (inside 50 mL falcon tubes) were placed in the freezer upon the moment of homogenization.
• Cut frozen lungs into small pieces (approximately 1 cm cubed).
• Homogenize your sample in a biosafety cabinet by grinding with a mortar and pestle to achieve a powdery consistency. Use liquid nitrogen to keep the samples frozen while being homogenized. Tip: separate your cubed sample before adding liquid nitrogen to prevent them from sealing together when freezing.
• Place portions of the tissue homogenates (0.25 g) inside sterile 2.0-ml screw-cap tubes for DNA extraction.
• Extract DNA from the tissue homogenates using a DNeasy PowerSoil kit (Qiagen Sciences Inc., Germantown, MD) according to the manufacturer’s protocol. For the last step, use 150 ul of elution buffer.
• Store extracted DNA at -20ºC before quality control.
• Use a nanodrop to determine the concentration of your DNA.
• Carry out a PCR using V3 / V4 primers to test whether there is bacterial DNA that can be amplified. For a detailed description, check out the PCR quality control protocol below.
• Send DNA samples (on dry ice) for 16S rRNA NGS after confirming that microbial DNA can be amplified.

Figure 11. Overview of the steps taken in the DNA extraction described in the ‘Microbiome DNA extraction protocol (homogenization)’.

Due to time constraints we were not able to optimize our previous protocol. In order to still gain results in time for the deadline, we used the following protocol to amplify bacteria in our lung sample using a growth chamber. We added a growth medium to our lung samples and incubated them in order to increase the number of bacteria present. Medium was extracted and analyzed further. This technique is only suitable to identify specific species in your sample, but the NGS cannot say anything about bacterial composition in percentages as you used a growth medium to amplify the number of bacteria in your sample.

Materials:

• Chicken lungs (acquired as a by-product from the meat industry).
• LB medium (steps to make this are described in growth media protocols).
• Biosafety cabinet.
• DNeasy PowerSoil Pro Kit

Protocol:

• Lungs (inside 50 mL falcon tubes) were placed in the freezer upon the moment of adding growth medium.
• Add 20 mL of LB medium to the falcon tubes containing the lungs. Don’t forget to create a negative control using the same medium.
• Thoroughly thaw the sample, and vortex for 2 minutes at max speed.
• Place the samples in an incubator at 37ºC and grow overnight.
• Vortex tubes for 2 minutes on max speed.
• Take out 250 ul of medium containing all of the grown bacteria present in the lung. If your sample somehow does not appear to have grown, put the tube back in the incubator overnight.
• Add the 250ul of medium containing the bacteria inside sterile 2.0-ml screw-cap tubes for DNA extraction.
• Extract DNA from the tissue homogenates using a DNeasy PowerSoil kit (Qiagen Sciences Inc., Germantown, MD) according to the manufacturer’s protocol. For the last step, use 150 ul of elution buffer.
• Store extracted DNA at -20ºC before quality control.
• Use a nanodrop to determine the concentration of your DNA.
• Carry out a PCR using V3 / V4 primers to test whether there is bacterial DNA that can be amplified. For a detailed description, check out the PCR quality control protocol below.
• Send DNA samples (on dry ice) for 16S rRNA NGS after confirming that microbial DNA can be amplified.

This protocol shows how you can use V3 / V4 primers from the hypervariable region of the 16S rRNA to check whether you have bacterial DNA in your sample that can be amplified.

Materials:

• DNA sample.
• Positive control (any bacterial DNA sample).
• Primers: target region V3 / V4 region 16S rRNA.
• Forward primer 338F sequences: 5'- ACTCCTACGGGAGGCAGCA-3'.
• Reverse primer 806R sequences: 5'- GGACTACHVGGGTWTCTAAT-3’.
• Phire Tissue Direct PCR Master Mix (Thermo Scientific).

Protocol:

• Prepare primers according to manufacturer instructions.
• Prepare PCR mixtures (total 10 ul):
• 5 ul phire tissue direct PCR master mix.
• 1 ul template DNA (your sample).
• 1 ul forward primer.
• 1 ul reverse primer.
• 2 ul MilliQ.
• Start a PCR on the following settings: one cycle of 95°C for 5 min, followed by 35 cycles of 98°C for 20 s, 50°C for 15 s, and 72°C for 1 min.
• Visualize PCR products using the agarose gel electrophoresis protocol described earlier.
• Identify whether you find bands around the 450 bp mark (expected size for the V3/V4 region).