E.coli culture

• Prepare LB medium by weighing and adding 25g of ready to mix LB agar mixture to 1L water in a sterile flask.
• Autoclave the broth and cool to room temperature.
• In a laminar flow chamber, transfer approximately 1 mL of overnight E. coli culture to the flask.
• Seal the mouth of the flask with sterile cotton (non-absorbant) plugs; ensure the flask is not tightly sealed.
• Incubate overnight at 37°C with continuous shaking (at 180 rpm).

Plating E. coli

• Prepare LB agar by weighing ready to mix LB broth, agar (in 2:1 ratio) and water to a sterile flask.
• Autoclave the medium and cool just enough to be able to handle the flask .
• In a laminar flow chamber, pour approximately 25-30 mL of the LB agar into sterile plate
• Allow the plates to set in the laminar flow chamber with lids slightly opened (The plates may also be stored inverted at 4°C for future use)
• Pick up E. coli colony from a plate with culture with a sterile inoculating loop.
• Streak the loop across the LB agar plate.

OMV isolation

• 250ml bacterial secondary culture and 4-6 autoclaved Oakridge tubes are placed inside the laminar hood.
• 250 ml secondary culture is poured into 4-6 Oakridge tubes equally.
• 4-6 Oakridge tubes are centrifuged at 8000 rpm at 4℃ for 30 min to pellet down the bacteria.
• Membrane filtration is done for the supernatant with 0.22 micron filter
• After that supernatant is collected into an autoclaved conical flask for ultracentrifugation.
• Polycarbonate tubes: not autoclavable, have to wash only with distilled water.
• Supernatants have to pour up to the mark ( aprox 90%of the tub, remaining 10% to grip the tube and to make sure that supernatant does not fall. Less than 90% will result in breaking of the tubes in ultracentrifuge)
• First take a beaker in the weighing machine and tare the weight.
• Next, put the beckman coulter tube’s cap into the weighing machine on the side.
• Supernatant is filled into the polycarbonate tubes up to the mark and then put into the beckman coulter tubes.
• Beckman Coulter tubes with polycarbonate tubes are placed into the center of the machine while a cap is also present into the machine and weight is taken.
• This procedure is repeated for all tubes and adjusted until the weight of all tubes become equal up to two decimal points.
• After that the tubes are weighed again for three times to recheck whether the weight is the same.
• Ultracentrifuge: 25000rpm, 4℃, 4 hours
• After 4 hours, let the rpm speed keep reducing, there would be a sound.
• Then, touch the vacuum button in screen, the vacuum would increase >1000 um
• Once it's done there would be another confirmation sound played.
• Next, gently slide open the lid and take out the rotor with the cassettes attached.
• Discard the supernatant from each of the 6 tubes.
• Add 0.5-1 ml of 1X PBS buffer to one of the tubes and pipette up and down in order to dissolve the OMVs into the buffer.
• Pipette out the dissolved OMV + buffer and pour into the next tube. Try shooting the PBS buffer in the place where OMVs have aggregated.
• Then collect the OMV-dissolved PBS buffer into the MCT tubes (1.5-2 ml)
• Again, rinse with 0.5-1 ml of 1X PBS buffer and collect in another MCT.
• Store the MCTs at -20℃.

OMV Purification

• Thaw the stored isolated OMV in MCTs on ice.
• Polycarbonate tubes: not autoclavable, have to wash only with distilled water and keep under UV for 5mins.
• Then take out 3ml OMVs from one MCT into the Polycarbonate tubes. Fill all the Polycarbonate tubes in this manner.
• First take a beaker in the weighing machine and tare the weight.
• Next, put the beckman coulter tube’s cap into the weighing machine on the side.
• The mixture is filled into the polycarbonate tubes up to the mark and then put into the beckman coulter tubes.
• Beckman Coulter tubes with polycarbonate tubes are placed into the center of the machine while a cap is also present into the machine and weight is taken.
• This procedure is repeated for all tubes and adjusted until the weight of all tubes become equal up to two decimal points.
• After that the tubes are weighed for three times to recheck whether the weight is the same.
• Ultracentrifuge: 30,000rpm, 4℃, 3hours
• After 3 hours, let the rpm speed keep reducing, there would be a sound.
• Then, touch the vacuum button in screen, the vacuum would increase >1000 um
• Once it's done there would be another confirmation sound played.
• Next, gently slide open the lid and take out the rotor with the cassettes attached.
• Discard the supernatant from each of the 6 tubes.
• Add 0.5-1 ml of 1X PBS buffer to one of the tubes and pipette up and down in order to dissolve the OMVs into the buffer.
• Then filter the OMV-dissolved PBS buffer with syringe filter into the MCT tubes (1.5-2 ml)
• Again, rinse with 0.5-1 ml of 1X PBS buffer, filter and collect in another MCT.
• Store the purified MCTs at -20℃.

1. DLS (Dynamic light scattering):

Sample Preparation:

• A diluted sample of OMV in MilliQ water (1:100 Dilution) is to be prepared.
• It is to be kept for sonication for 15 minutes
• The cuvette is to be cleaned with the respective solvent and washed two-three times.
• The experiment has to be started with 10ul of sample in the cuvette.
• PBS buffer has to be added till the required height in the cuvette.
• The cuvette has to be placed in the proper orientation as shown in the instrument.

Measuring the size:

 The instrument documentation is attached here.
 Follow the steps mentioned above.
 The Parameters that need to be set are as follows:
 a. Manual Measurement
 b. Measurement Type: Size
 c. Sample name: iGEM_OMV_(SAMPLE NAME -S1)_(DD/MM)
 d. Dispersant - MilliQ Water
 e. Temperature- Standard Mention equilibrium time 60 second.
 f. Select Cell - Select the cuvette we will be using.
 g. Measurement Criteria.
 i. Number of Runs - 15
•ii. Run Duration - 10
•iii. Number of measurements per scan - 5

Zeta Charge Measurement:

 During Sample Preparation use MILLIQ WATER as your solvent.
 Follow the above given steps for SIZE measurement instead of selecting measurement type Size, select ZETA.
 Materials Required:
 a. MilliQ Water
 b. PBS Wash buffer
 c. Pipette 100µl,10µl,1000µl.
 d. Cuvette
 e. Tissue Paper
 f. Sonicator

2. Bradford Assay:

Materials required:

• BSA
• Bradford reagent
• 96 well plate
• 2.5ul, 10ul, 20ul, 100ul and 200 ul pipettes
• Aluminum foil
• Distilled water
• Spectrophotometer

Standard Curve preparation:

• Prepare 1 ml of 3mg/ml stock solution of BSA in MCT.
• From stock-solution, prepare 100ul of BSA of the following concentrations-0.05ug/ul, 0.1ug/ul, 0.5ug/ul, 0.8ug/ul, 1ug/ul and 1.2ug/ul- each in a 1.5ml MCT.
• For each concentration, take 10 ul of BSA in 3 consecutive wells in a 96 well plate, then add 190 ul of Bradford reagent in each well.
• Wrap the entire plate in aluminum foil and keep the plate for 15 minutes.
• After 15 minutes, measure OD595.
• Prepare a standard curve using the data obtained. Protein content should be on x-axis and OD value should be on y-axis.

Protein Quantification:

• In a 96 well plate, add 10 uL of OMV sample to each well.
• Add 190 uL of Bradford reagent.
• Cover the plate with aluminum foil, incubate in the dark for 15 minutes.
• Measure the OD595 nm. Using the standard curve constructed, determine the protein concentrations.

3. SEM (Scanning Electron Microscopy):

• The cover slip is broken into small pieces so that it can fit into the SEM holder.
• 10-15 ul of concentrated OMV solution is to be put onto the cover slip and waited for it to dry.This step was done in the laminar hood.
• A dish or well is to be taken where the cover slip has to be kept and then covered with 2.5 % (v/v) glutaraldehyde (prepared in PBS; pH 7.4) at room temperature (RT) overnight in the dark to fix the specimens.
• Fixed samples are to be washed thrice with PBS for 10 min each followed by sequential dehydration in 35 %, 50 %, 70 %, 95 % ethanol for 10 min each and 100 % ethanol for 1 h for complete dehydration.
• Finally, fixed and dehydrated samples are to be vacuum-dried overnight.
• Specimens are to be sputter-coated with gold and examined using a Supra 55 Carl Zeiss scanning electron microscope.

4. TEM (Tunelling Electron Microscopy):

Preparation of 2% Uranyl acetate(UA)

• 200mg of UA is to be weighed
• 1ml of MilliQ Water is to be added

Note/Precautions:

• UA is a light sensitive compound so experiments need to be performed in low light areas to avoid any imaging issues.
• It is a radioactive hazardous compound

Sample Preparation:

 a. A blotting paper is to be placed on the petri dish so that it covers all the transparent portion.  b. Forceps are to be carefully held at the edge of the TEM Grid mid air.  c. 15ul of sample is to be added and kept for 1 minute  d. 15ul of 4%UA is to be added and kept for 4 minutes.  e. Excess sample is to be slowly cleared out from the edges of the TEM grid using blotting paper.  f. The grids are to be placed on your petri dish and desiccated.

Xanthomonas culture:

DSF isolation:

1. Inoculate XooBXO43 in primary culture with antibiotics (chloramphenicol).
2. Grow to saturation at 28*C.
3. Inoculate for secondary culture in media without antibiotics.
4. Grow to saturation at 28*C.
5. Pellet the cells at 8000rpm for 10 minutes at room temperature.
6. Collect the supernatant.
7. Add water-saturated ethyl acetate to the supernatant in 5:2 ratio.
8. Stir with magnetic beads for at least 45-60 minutes.
9. Centrifuge at 8000 rpm for 10 minutes at room temperature.
10. Take the upper layer and dry with speed vacuum / air dry at 37*C.
11. Dissolve the extracted DSF in methanol, i.e., 250ul of methanol in need for 250ml.

Flow Cytometry sample preparation and Confocal Microscopy Sample Preparation Verification of OMVs Fusion with Xanthomonas:

1. 200ul of OMV sample in PBS buffer are labelled with 1 ug/ml dye.
2. Then the OMVs are washed in PBS by ultracentrifugation _rpm,_hrs.
3. Xoo Cells are adjusted to OD600 = 1.0 in PBS buffer.
4. The Xoo cell cultures are further divided into two different sets.
5. First one will be a control set and the second one will be fusion set.
6. For fusion set:
 a. OMVs Labelled with 5ul dye to be incubated with Xoo in an eppendorf tube/Facs tube for 30 minutes at R/T.
 b. Different time points of incubation can be applied. 7. Two different measurements of Mean Fluorescent intensities will be compared between both the sets.
4

Alternative protocol:  For fusion set:  a. Label the OMVs with 1 ug/ml of Rhodamine B dye
 b. Label the bacterial cells 1 ug/ml of FM1-43 dye.
 c. Wash the bacterial cells with PBS 3 times.
 d. Incubate the bacterial cells with OMV for 30 minutes at Room Temperature.
 e. MFI changes are calibrated.  For confocal:  1. DAPI mounting dye was used in 1ug/ml concentration to mount the sample in the slides, the incubation mixture was mounted.
2. Similar protocol was applied for FM1-43 mounting dye.

Chemically competent E.coli preparation (for Top10F):

1. We have to recover the cells by centrifugation at 4100rpm or 2700 × g for 10min at 4˚C.
2. We need to decant the supernatant and then stand the tubes in an inverted position on a pad of paper towel for 1min to allow the last traces media to drain away.
3. We have to add 15ml of ice-cold MgCl2-CaCl2 solution (80mM MgCl2, 20mM CaCl2) to the pellet. Then resuspend each pellet by swirling or gentle vortex as the cells can burst due to hash vortexing (The cells shouldn’t have clumps, it should mix well).This has to be done in cold(4°C) and aseptic conditions. [CaCl2 can alone produce competent cell. But we are increasing its competency by adding one more positive ion.Here MgCl2 used is 4times more than CaCl2, so that in cell wall maximum MgCl2 is there.]
4. We have to again recover the cells by centrifugation at 4100rpm or 2700 × g for 10min at 4˚C.
5. Now we have to decant the medium from the cell pellets and stand the tubes in an inverted position on a pad of paper towel for 1-2min to allow the last traces media to drain away
6. We have to add 2ml of ice-cold 0.1M CaCl2 solution to the pellet for each 50ml of original culture. Now resuspend each pellet by swirling or gentle vortex (no clumps should be present). This should be done in aseptic conditions. [This time CaCl2 is maximum as it will rotate , centrifuge so in that case few CaCl2 is left behind].
7. Before freezing we have to add 70μl of DMSO(anti-freezing agent is toxic) drop by drop in each 2ml of CaCl2 solution mixed with pellet,otherwise, there is a high chance that cell will die if the total concentration of DMSO is added at once. We need to shake the tube after adding each drop so that it mixes well.
8. We need to keep in ice for 15min so that it mixes well.
9. Again we have to add 70μl of DMSO drop by drop and shake and swirl for some time. [DMSO is a toxic agent, so for actual content of 2ml we should add around 150µl. But if we give it at a time, then it could be harsh for the cells. So we let the cells settle down and then add the next round of DMSO so that they all over spread to the media.]
10. We have to aliquot the competent of 50-60μl in 0.5 ml micro centrifuge tube.
11. Now we have to Flash freeze(provides inter-cellular conditions) in liquid nitrogen(-196°C) and store it in -80˚C. [This is done to prevent cells from dying]

Electrocompetent E.coli preparation:

1. Grow an overnight culture of each strain in LB medium.
2. Prepare 10 ml of fresh LB medium in a 50 ml flask for each strain.
3. Inoculate with 100 μl of the overnight, stationary-phase culture.
4. Grow the cells for approximately 2-3 hours, until they reach mid-exponential phase (In general, this is an OD600 of ~0.6).
5. Transfer the cells to 15 ml Falcon conical tubes.
6. Pellet the cells by centrifugation for 5 minutes at 6,000 RPM. Remove promptly and pour off supernatant.
7. Wash by adding 10 ml of chilled 10% glycerol to each tube, then vortexing vigorously to resuspend the pellet. Centrifuge for 3.5 minutes. Remove promptly and pour off supernatant. Repeat for at least four wash cycles in 10% glycerol.
8. Resuspend in approximately 100 μl of 10% glycerol to make a 100x concentration of the initial culture.
9. Divide into 30-50 μl aliquots in 0.5 or 1.7 ml tubes. Freeze or proceed directly to electroporation.

Chemical Transformation in E.coli 10-beta:

1. Thaw a tube of NEB 10-beta competent E.Coli cells on ice until the last ice crystal disappears. Mix gently and carefully pipette 50ul of cells into a transformation tube on ice.
2. Add 1-5ul containing 1pg-100 ng of plasmid DNA to the cell mixture. Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.
3. Place the mixture on ice for 30 mins. Do not mix.
4. Heat shock at exactly 42°C for exactly 30secs. Do not mix.
5. Place on ice for 5 mins. Do not mix.
6. Pipette 950 ul of room temperature NEB 10-beta/SOC into the mixture.
7. Place at 37°C for 1 hour. Shake vigorously (250rpm) or rotate.
8. Warm selection plates to 37°C.
9. Spread 50-100ul from the incubated bacterial cells in SOC onto a selection plate and incubate overnight at 37°C.

Electrotransformation in E.coli 10-beta:

1. Thaw the electrocompetent cells on ice.
2. To the electrocompetent cells, add 1-3 μl of DNA (lesser than 100 ng of DNA).
3. Mix by gently flicking the tube containing the electrocompetent cell + DNA mixture. Let the mixture sit on ice for 1-10 minutes.
4. Pipette the mixture into a chilled cuvette, making sure that the mixture is at the bottom of the cuvette by gently tapping the cuvette on a flat surface (Be sure to wipe any condensation off the sides of the cuvette before electroporation).
5. Place the cuvette in the pulser and press the "Pulse" button (For the TOP10F Electrocompetent E. coli cells, the Ec l setting is fine)
6. After electroporation, add 500-1000 μl of SOC/LB to the cuvette to recover the cells.
7. Transfer the mixture to a 1.5 mL microcentrifuge tube.
8. Incubate for ~30-60 minutes at 37°C or other appropriate temperature in a shaking incubator (Be sure to place the tube on its side so the transformed cells will grow properly).
9. Plate the cells (~ 50 μl) on an LB plate containing the appropriate antibiotic.
10. Incubate overnight at 37°C or other appropriate temperature.

Electrocompetent Xanthomonas preparation:

1. Give 3mL Xanthomonas primary culture in 2.5% LB, incubate for 16-20 hours at 28°C and 250rpm.
2. Give 30mL secondary culture in 2.5% LB with 300uL inoculum, incubate at 28°C for 3.5hours at 250rpm.
3. Transfer the culture to an Oakridge tube and centrifuge at 6000rpm for 5minutes at 4°C. Discard supernatant and resuspend pellet with 30mL dH2O.
4. Centrifuge at 6000rpm for 8minutes at 4°C. Discard supernatant and resuspend pellet in 15mL dH2O.
5. Centrifuge at 6000rpm for 8minutes at 4°C. Discard supernatant and resuspend pellet in 5mL dH2O.
6. Centrifuge at 6000rpm for 8minutes at 4°C. Discard supernatant and resuspend pellet in 100uL dH2O. Entire pellet will not dissolve, dissolve as much as possible and transfer 100uL of cell culture to a 1.5mL MCT placed in ice bucket.
7. Transfer 100ng of plasmid to the cell culture and keep for 15minutes. Meanwhile, cool the electrocuvette at 4°C.
8. After 15minutes, transfer 100uL of the mixture to the electrocuvette and again keep in 4°C for 15minutes.
9. Electrocute the mixture inside a cuvette using a pulse voltage.
10. Quickly add 1mL of LB inside the electrocuvette and mix well via pipetting.
11. Transfer 1mL of the culture to a fresh 1.5mL autoclaved MCT. Incubate it for 3.5hours at 28°C and 250rpm.
12. Centrifuge the solution at 6000rpm for 5minutes. Discard supernatant and resuspend pellet in 100uL LB.
13. Spread the 100uL culture on an antibiotic LB agar plate and incubate for 1-2 days at 28°C.

Plasmid isolation protocol:

1. Pellet 1-5ml ( do not exceed 15 OD units) Bacterial culture by centrifugation for 30secs. Discard the supernatant.
2. Resuspend pellet in 200ul Plasmid Resuspension Buffer. Vortex or pipette to ensure cells are completely resuspended. There should be no visible clumps.
3. Add 200ul Plasmid Lysis Buffer, gently invert the tube 5-6times, and incubate at room temperature for 1minute. The color should change to dark pink and the solution will become dark and viscous. Do not vortex.
4. Add 400ul of Plasmid Neutralization Buffer, gently invert tube until neutralized and incubate at room temperature for 2 minutes.The Sample is neutralized when color is uniformly yellow and precipitate forms. Do not vortex.
5. Centrifuge Lysate for 2-5minutes. For best results and especially for culture volumes >1ml, a 5min spin ensures efficient RNA removal by RNase A.Pellet should be compact, spin longer if needed.
6. Carefully transfer supernatant to the spin column and centrifuge for 1min. 9Discard the flow through.
7. Re-insert column in the collection tube and add 200ul of Plasmid Wash Buffer 1. Centrifuge for 1min. Discarding the flow through is optional.
8. Add 400ul of Plasmid WSh Buffer 2 and Centrifuge for 1min.
9. Transfer column to a clean 1.5ml MCT. Carefully ensure that the tip of the column does not come in contact with the flow through. If there is any doubt, re-spin the column for 1min.
10. Add >= 30ul DNA Elution Buffer to the center of the matrix. Wait for 1min, then spin for 1min to elute DNA. Nucleus free water can also be used to elute the DNA. Yield may slightly increase if a larger volume of DNA Elution Buffer is used, but the DNA will be less concentrated. For larger size DNA, (>= 10kb), heating the Elution Buffer to 50°C prior to use can improve yield.

Gel extraction:

1. Excise the DNA fragment from the agarose gel with a clean, sharp scalpel. Minimize the size of the gel slice by removing extra agarose.
2. Weigh the gel slice in a colorless tube. Add 3 volumes of Buffer QG to 1 volume of gel (100 mg ~ 100 µl). For example, add 300 µl of Buffer QG to each 100 mg of gel. For >2% agarose gels, add 6 volumes of Buffer QG. The maximum amount of gel slice per QIAquick column is 400 mg; for gel slices >400 mg use more than one QIAquick column.
3. Incubate at 50°C for 10 min (or until the gel slice has completely dissolved). To help dissolve gel, mix by vortexing the tube every 2–3 min during the incubation.
IMPORTANT: Solubilize agarose completely. For >2% gels, increase incubation time.
4. After the gel slice has dissolved completely, check that the color of the mixture is yellow (similar to Buffer QG without dissolved agarose). If the color of the mixture is orange or violet, add 10 µl of 3 M sodium acetate, pH 5.0, and mix. The color of the mixture will turn to yellow. The adsorption of DNA to the QIAquick membrane is efficient only at pH ≤7.5. Buffer QG contains a pH indicator which is yellow at pH ≤7.5 and orange or violet at higher pH, allowing easy determination of the optimal pH for DNA binding.
5. Add 1 gel volume of isopropanol to the sample and mix. For example, if the agarose gel slice is 100 mg, add 100 µl isopropanol. This step increases the yield of DNA fragments <500 bp and >4 kb. For DNA fragments between 500 bp and 4 kb, addition of isopropanol has no effect on yield. Do not centrifuge the sample at this stage.
6. Place a QIAquick spin column in a provided 2 ml collection tube.
7. To bind DNA, apply the sample to the QIAquick column, and centrifuge for 1 min. The maximum volume of the column reservoir is 800 µl. For sample volumes of more than 800 µl, simply load and spin again.
8. Discard flow-through and place QIAquick column back in the same collection tube. Collection tubes are re-used to reduce plastic waste.
9. (Optional): Add 0.5 ml of Buffer QG to QIAquick column and centrifuge for 1 min. This step will remove all traces of agarose. It is only required when the DNA will subsequently be used for direct sequencing, in vitro transcription or microinjection.
10. To wash, add 0.75 ml of Buffer PE to QIAquick column and centrifuge for 1 min. Note: If the DNA will be used for salt sensitive applications, such as blunt-end ligation and direct sequencing, let the column stand 2–5 min after addition of Buffer PE, before centrifuging.
11. Discard the flow-through and centrifuge the QIAquick column for an additional 1 min at ≥10,000 x g (~13,000 rpm).
IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation.
12. Place QIAquick column into a clean 1.5 ml microcentrifuge tube.
13. To elute DNA, add 50 µl of Buffer EB (10 mM Tris·Cl, pH 8.5) or ddH2O to the center of the QIAquick membrane and centrifuge the column for 1 min at maximum speed. Alternatively, for increased DNA concentration, add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge for 1 min.

Gibson Assembly:

Set up the following reaction on ice:
1. Incubate samples in a thermocycler at 50°C for 15 minutes (when 2 or 3 fragments are being assembled) or 60 minutes (when 4–6 fragments are being assembled). Following incubation, store samples on ice or at –20°C for subsequent transformation.
Note: Extended incubation up to 60 minutes may help to improve assembly efficiency in some cases
2. Transform competent E. coli cells with 2 μl of the chilled assembled product, following the transformation protocol.

Colony PCR:

1. Pick a single colony using a sterile pipette tip with a pipette set to 5µl and pipette up and down to mix in 25µl of water in a PCR tube. Then transfer 5 µl of this into your empty PCR tube. Do this for about 6 colonies. Save the remaining 20µl for later.
2. The PCR reactions are set up as below. Because you are doing this reaction multiple times, you should make a “Master Mix” of PCR components so you don’t have to individually pipette them into tubes every time. This becomes especially time saving when you do this for 10 or more reactions. Make your Master Mix as enough reagent for one more reaction than you are doing, e.g. if you are PCRing 6 colonies, make Master Mix for 7 colonies.

3. To your tubes containing the 5µl of bacterial cells, add 15µl of your 7x Master Mix. Vortex to mix and spin down in the capsule microfuge.
4. Put your PCR strips in the thermocycler for the following cycle (probably saved as some file called “GoTaq”). 95 5 min 30x: 95 45 s 55 30 s 72 1 min per kb 72 10 min
5. Analyze products on a 1% agarose gel. The GoTaq mix already has gel loading dye in it, so you can just directly load 5-10 µl of your PCR reaction into a gel.
6. For all positive bands on the gel, take the rest of the bacterial cells from step 1 and inoculate them into an overnight LB (+antibiotic) for miniprep the next day.