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Imperial iGEM 2014

Here you can find the protocols we used throughout the summer

G. xylinus Protocols

G. xylinus HS media and culturing

  • Materials
    • For 500ml media:
    • 10g glucose (2%w/v)
    • 2.5g yeast extract (0.5% w/v)
    • 2.5g peptone (0.5% w/v)
    • 1.35g Na2HPO4 (0.27% w/v)
    • 0.75g citric acid (0.15% w/v)
    • 500ml distilled H20
    • 0.5mL cellulase if HS+cellulase media required
    • 7.5g of agar (agar-agar) if making HS-agar plates
    • (antibiotics as necessary)
  1. Add 250ml dH20 to glucose in one bottle and 250ml dH20 to the rest in a second bottle. In incompletely distilled water, glucose will form a solid mass, so stir vigorously immediately after adding water. Autoclave both bottles to sterilize media and pour glucose solution in sterile conditions (next to a Bunsen burner or in a flow hood) into the second bottle. Autoclaving glucose separately from amino acids avoids Maillard reaction, which can result in the formation of toxic byproducts in the media.
  2. Streak/inoculate Gluconacetobacter onto plates or into media.
  3. Incubate plates at 30°C inverted. Colonies will appear in 48-72 hours.
  4. Incubate liquid HS-cultures at 30 °C standing. Standing culturing results in low growth rate, but avoids the formation of cellulose non-producing mutants, which have been reported to appear in shaking conditions.
  5. For quick growth, grow with shaking at 180rpm, 30C, with the addition of cellulase. NB! This selects for cellulose non-producing mutants. They can be identified as smooth colonies on plates (cellulose producing colonies have a rough colony morphology - see section Gluconacetobacter for images).

Preparing electrocompetent G. xylinus cells

  • Materials
    • HS+cellulase media
    • Ice bucket and ice
    • Temperature controlled centrifuge
    • 1mM HEPES (ph7.0) 80mL
    • 15% glycerol
    • 50 mL tubes
    • P1000 pipette and tips
    • Stripettes and automatic pipette for larger volumes
    • Shaker at 30 °C, 180rpm
  1. If the goal of transformation is to produce cellulose-producing transformed G.xylinus, use regular HS media for culturing. If cellulose production after transformation is not primary and speed is required, HS-cellulase medium can be used. Usage of HS-cellulase medium during growth results in the formation of a higher number of cellulose negative mutants, but results in much higher transformation efficiencies, and requires less time. Even with HS-cellulose medium, cellulose producing colonies can be identified on the plate after transformation, as cellulose-producing colonies differ in morphology from cellulose non-producing colonies (see section Gluconacetobacter for images of colony morphology).
  2. Inoculate 5ml of HS or HS+cellulase medium with Gluconacetobacter.
  3. Incubate at 30°C, 180 rpm shaking overnight.
  4. Next day, pour 30mL of HS or 15ml of HS+cellulase medium into each of four 50mL tubes. If using regular HS medium, vortex the tubes for 3 minutes to release cells from the cellulose pellicle
  5. To each tube, add 1mL of overnight culture and incubate with shaking at 180rpm, 30C.
  6. Incubate overnight or until OD600 of around 0.4-0.7 is reached. When using HS-cellulase media, OD600 can reach up to 0.6 -0.7, however when using regular HS medium, OD600 measurement is disturbed by the cellulose pellicle, and can reach up to 0.2. Vortex tubes for 3 minutes before taking the measurements.
  7. If using regular HS medium, add 62ul (0.2% v/v) Celluclast cellulase to each tube and incubate at 30C, 180rpm shaking for 2 hours, or until the cellulose pellicle is degraded to completely release the cells. This does not result in higher formation of cel- mutants, as short incubation time and nutrient depletion does not allow for proliferation of cel- mutants.
  8. Before continuing, set up the necessary materials:
    1. Pre-cool centrifuge to 4°C
    2. Prepare ice bucket
    3. Chill 1mM HEPES buffer and 15% glycerol buffer on ice
  9. Once the cultures reach desired OD600, take them out of incubation and put them on ice for 10 minutes (the tube should feel cool).
  10. From here on, keep cells always cool, at or below 4°C
  11. After cooling, spin the tubes in a refrigerated (4°C) centrifuge for 12min at 4100rpm.
  12. Pour off supernatant carefully, taking care not to pour off the pellet. G. xylinus does not pellet as easily as E.coli, most likely due to the buffering effects of cellulose. If the pellet is not attached to the wall after centrifugation, smear the pellet onto the wall of the tube and centrifuge again using longer centrifugation times. Re-suspend bacteria in 10mL HEPES: re-suspend first using 1ml HEPES and a P1000 pipette, then add 9ml of HEPES using a stripette; it is much easier to re-suspend the pellet fully using a P1000. Do not use a vortexer.
  13. Pool the samples into a single 50ml tube.
  14. Centrifuge again for 14 minutes at 4100 rpm and 4 °C temp. Use a balancer tube with 40ml water.
  15. Pour off supernatant again, re-suspend pellet in 10mL ice-cold HEPES on ice as before.
  16. Centrifuge again for 14 minutes at 4100 rpm, 4 °C.
  17. Pour off supernatant and re-suspend pellet in 4mL ice cold 15% glycerol solution.
  18. Pipette 50ul aliquots into tubes. Label tubes properly.
  19. Store samples on ice for immediate use or freeze 50ul aliquots in-80°C. According to some reports, the efficiency of electrocompetent cells reduces after each freezing, so immediate use may result in highest efficiencies.

Quantification of cellulose production

  1. Add 50ml of HS medium (or other medium of choice) to 250ml conical flask
  2. Grow G.xylinus in HS medium for 7 days standing, at 30C. Don’t seal the flasks hermetically in order to allow diffusion of oxygen (seal using foam buns)
  3. After 7 days of growth, wash the cellulose twice with distilled water
  4. Add 50ml of 0.1M NaOH to cellulose, incubate at 65C for 2 hours
  5. Wash the cellulose twice using distilled water
  6. Place the formed cellulose pellicle on baking paper and dry the pellicle at 65 degrees 2 hours-overnight. Move the pellicle in the meantime, in order to avoid it sticking to the paper, which can skew your results.
  7. Place the pellicle into a vacuum desiccator for 2 hours
  8. Weigh the pellicle using a high-sensitivity scale.

Transformation of G.xylinus using electroporation

  • Prepare the following
    • Plasmid DNA
    • Electrocompetent G.xylinus cells (in 50µl or 100µl aliquots)
    • Ice bucket and ice
    • 1mm path electrocuvettes
    • 1.5ml microcentrifuge tubes or PCR tubes
    • An electroporator - set at 2.5kV and 5.9ms
    • HS+cellulase medium
    • HS agar plates with appropriate antibiotic
    • One aliquot of electrocompetent cells for positive control and one aliquot for negative control.
    • If a plasmid known to work in G. xylinus exists, use this as positive control .
  1. Set up the electroporator with correct settings: at 2.5kV, 6-8ms, 400ohm resistance, 25microF capacitance.
  2. Prepare 800µL HS+cellulase media containing 1.5ml culture tubes and pre-heat to 30 °C. Prepare SOC medium at 37°C. Prepare a space for shaking.
  3. Prepare ice bucket, place plasmid DNA and electrocuvettes on ice and thaw electrocompetent cells on ice. NB! Make sure DNA is desalinated before use – ionic solution can otherwise cause arcing.
  4. Add 2 µL of plasmid DNA to 100 µL of concentrated cells in a cold microcentrifuge or PCR tube and mix well by pipetting. Add plasmid DNA also to one aliquot of electrocompetent E.coli cells (positive control; alternatively also add another known plasmid as pos. control). NB! Do not add plasmid DNA to one aliquot of electrocompetent G. xylinus (negative control)
  5. Transfer the cell/DNA mixture and positive and negative controls to a cold 0.2-cm electroporation cuvette. Dry any water condensate outside of the cuvette (using labcoat), place the cuvette into the electroporator, and apply the pulse.
  6. Transfer the pulsed cells into 800 µL of HS+cellulase medium in a culture tube. Transfer E. coli into SOC medium. NB! Prepare everything beforehand and do it quickly.
  7. Incubate the culture tubes with shaking (170 rpm) at 30 °C overnight

gDNA Library preparation for genome sequencing using Illumina Nextera kit

This is a modified protocol for using Illumina Nextera kit for preparation of gDNA library for genome sequencing. We have modified the original Illumina protocol to be amenable for low sample number and have a lower cost. We used this protocol to prepare gDNA libraries of G. xylinus ATCC 53582 strain and the G. xylinus strain we isolated from Kombucha tea at the beginning of this summer. For the full protocol and required materials, see Illumina Nextera Kit user guide. NOTE: When first learning about gDNA library preparation, we have found SeqAnswers forum highly informative.


  1. Prepare 200ul PCR tubes, ice bucket, ice
  2. Remove the TD, TDE1, and genomic DNA from -15° to -25°C storage and thaw on ice.
  3. Remove RSB from -15° to -25C storage and thaw at room temperature.
  4. After thawing, ensure all reagents are adequately mixed by gently inverting the tubes 3–5 times, followed by a brief spin in a microcentrifuge. NOTE: Ensure the reaction is assembled in the order described for optimal performance. Some sources recommend setting up the reaction on ice, as transposons may exhibit a low activity at room temperature
  5. Label the PCR tubes with a smudge resistant pen - important, as labels tend to degrade in a thermocycler.
  6. Add 20 μl of genomic DNA at 2.5 ng/μl (50 ng total) to each PCR tube
  7. Add 25 μl of TD Buffer to the wells containing genomic DNA. Change tips between samples. NOTE:Calculate the total volume of TD for all reactions, and divide among an appropriate number of tubes in an 8-well PCR strip tube.
  8. Add 5 μl of TDE1 to the tubes containing genomic DNA
  9. Gently pipette up and down 10 times to mix the reaction.
  10. Centrifuge at 280g for 1 minute
  11. Place the PCR tubes into the thermocycler and run the following program:
    1. Heated lid on
    2. 55°C for 5 minutes
    3. Hold at 10°C
  12. While tagmentation is ongoing, label 1.5ml tubes accordingly for the next step and add 180ul of Zymo DNA binding buffer
  13. Also, remove NPM, PPC, and the index primers from -15° to -25°C storage and thaw on a bench at room temperature.
  14. Allow approximately 20 minutes to thaw NPM, PPC, and index primers
  15. After tagmentation reaction has finished, proceed immediately to Zymo cleanup - transport samples on ice and do not wait before proceeding to Zymo cleanup, as transposons remain active at lower temperatures, albeit at low level, which may result in overtagmentation and fragments shorter than optimal.

Zymo cleanup protocol

  1. Transfer 50 μl of tagmentation reaction into 1.5ml tubes containing Zymo binding buffer
  2. Gently pipette up and down 10x to mix well
  3. Transfer the solution to Zymo spin columns
  4. Centrifuge at 10000rpm for 1 minute.
  5. Discard the flow-through or the collecting tube and place the column into a new collection tube
  6. Wash the Zymo spin column twice by:
    1. Pipette 300ul of Zymo wash buffer to the tubes
    2. Centrifuge at 10000rpm for 1 minute, discarding the flow through
    3. Repeat the wash step for a total number of 2 washes.
    4. Centrifuge at 10000rpm 1 min to ensure no wash buffer remains.
    5. Add 25 μl of RSB directly to the column matrix in each well. Confirm visually that RSB has absorbed onto the matrix, and is not on the side of the tube. If it is, flick the tube gently to force the liquid onto the matrix
    6. Incubate the tubes for 5 minutes at room temperature. If time is not a limiting factor, increase incubation time to 15 minutes, as this may result in higher elution effieciencies
    7. Centrifuge the tubes at 10000rpm for 1 minute.
    8. Place the tubes on ice until proceeding to PCR


  1. After NPM, PPC, and Index primers are completely thawed, gently invert each tube 3–5 times to mix and briefly centrifuge the tubes in a microcentrifuge. Use 1.7 ml Eppendorf tubes as adapters for the microcentrifuge.
  2. Label PCR tubes according to your samples
  3. Add 5 μl of index 2 primers (white caps) to each tube respectively.
  4. Add 5 μl of index 1 primers (orange caps) to each tube respectively.
  5. Add 15 μl of NPM (Nextera PCR master mix) to each well of the NAP1 plate containing index primers.
  6. Add 5 μl PPC (PCR primer cocktail) to each well containing index primers and NPM.
  7. Transfer 20 μl of purified tagmented DNA to the corresponding PCR tube
  8. Gently pipette up and down 3–5 times to thoroughly combine the DNA with the PCR mix.
  9. Spin down the tubes with quick centrifugation .
  10. Ensure that the thermocycler lid is heated during the incubation.Run the PCR using the following program:
    1. 72°C for 3 minutes
    2. 98°C for 30 seconds
    3. 5 cycles of:
      1. 98°C for 10 seconds
      2. 63°C for 30 seconds
      3. 72°C for 3 minutes
    4. Hold at 10°C
  11. Ensure that the 72°C step preceeds the rest of the program.
  12. SAFE STOPPING POINT – can store the DNA in thermocycler overnight, or at 2-8°C for 2 days

Clean-up of PCR using AMPure beads

  1. Bring the AMPure XP beads to room temperature.
  2. Prepare fresh 80% ethanol from absolute ethanol. (Always prepare fresh 80% ethanol for wash steps. Ethanol can absorb water from the air impacting your results.)
  3. xCentrifuge the PCR tubes containing the limited cycle PCR product at quickly to spin down the liquid.
  4. Label new 1.5ml tubes according to the samples for purification steps
  5. Transfer 50 μl of the PCR product from the PCR tubes into new 1.5ml tubes. Change tips between samples.
  6. Vortex the AMPure XP beads for 30 seconds to ensure that the beads are evenly dispersed.
  7. Add 30 μl of AMPure XP beads to each tube containing the PCR product. For 2x250 runs on the MiSeq, add 25 μl of AMPure XP beads to each tube.
  8. Gently pipette mix up and down (gently, so as not to introduce bubbles) until solution is homogeneous. Make sure to pipette, no to vortex the solution, as this results in liquid collecting under the tube cap. This can't be removed easily as centrifugation also results in sedimentation of AMPure beads.
  9. Incubate the tubes (containing AMPure beads and PCR product) at room temperature without shaking for 5 minutes.
  10. Place the plate on a magnetic stand for 2 minutes or until the supernatant has cleared.
  11. With the tubes on the magnetic stand, carefully remove and discard the supernatant. Pieptte carefully, as AMPure beads may follow the surface of the liquid. If any beads are inadvertently aspirated into the tips, dispense the beads back to the plate and let the plate rest on the magnet for 2 minutes and confirm that the supernatant has cleared.
  12. With the tubes on the magnetic stand, wash the beads with freshly prepared 80% ethanol as follows:
    1. Add 200 μl of freshly prepared 80% ethanol to each sample well. You should not resuspend the beads at this time.
    2. Incubate the plate on the magnetic stand for 30 seconds or until the supernatant appears clear.
    3. Carefully remove and discard the supernatant.
  13. With the tubes on the magnetic stand, perform a second ethanol wash as follows:
    1. Add 200 μl of freshly prepared 80% ethanol to each sample.
    2. Incubate the plate on the magnetic stand for 30 seconds or until the supernatant appears clear.
    3. Carefully remove and discard the supernatant. Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol.
  14. With the tubes still on the magnetic stand, allow the beads to air-dry for 15 minutes.
  15. Remove the tubes from the magnetic stand. Add 32.5 μl of RSB to each well of the NAP2 plate.
  16. Gently pipette mix up and down until the solution is homogenous, changing tips after each column.
  17. Incubate at room temperature for 2 minutes.
  18. Place the tubes on the magnetic stand for 2 minutes or until the supernatant has cleared.
  19. Label new 1.5ml tubes accordingly
  20. Carefully transfer 30 μl of the supernatant into the new tubes. These will contain your purified library.
  21. Store the library in -20C until further processing.

General/E. coli Protocols

LB Broth Preparation

  1. Add 25g Luria Broth to 1L demineralised water
  2. Autoclave

LB Agar Preparation

  1. Add 25g Luria Broth and 15g Agar to 1L demineralised water
  2. Autoclave

Rubidium Chloride Competent Cells

  1. Inoculate 1ml of cell culture (grown overnight) into flask containing Psi Broth
  2. Incubate for 2h at 37°C
  3. Transfer culture into sterile falcon tube, place in ice for 15min
  4. Centrifuge at 4000rpm for 5 minutes
  5. Discard supernatant, then add TfBI buffer and resuspend pellet
  6. Place in ice for 15 minutes
  7. Centrifuge at 4000rpm for 5 minutes
  8. Discard supernatant, then add TfBII buffer and resuspend pellet
  9. Produce 50ul aliquots

Heat-Shock Transformation

  1. Add insert DNA to cell aliquot
  2. Place on ice for 15 minutes
  3. Heat-shock: place in a 42°C heat-block for 45 seconds
  4. Place samples back on ice for 2 minutes
  5. Add LB
  6. Incubate at 37°C for 30-60 minutes
  7. Centrifuge at 4000rpm for 5 minutes
  8. Discard 300uL LB
  9. Resuspend cells in remaining 200uL LB
  10. Plate out
  11. Incubate at 37°C overnight.

80% Glycerol Preparation

  1. Add 80ml 99.7% glycerol to 20ml demineralized water
  2. Autoclave

Glycerol Stock Preparation

  1. Cultures plated on LB Agar + antibiotic and grown at 37°C overnight.
  2. A 5ml LB culture in LB+antibiotic inoculated from a single, freshly growing colony.
  3. Cultivate for 16h at 37°C, with constant shaking
  4. 0.5ml of this culture inoculated into sterile vial
  5. Add 0.5ml of 80% glycerol
  6. Vortex
  7. Spin down
  8. Freeze them at -80 degrees

QIAprep Spin Miniprep Kit

  • Materials per sample
    • 250ul P1 buffer (suspension buffer)
    • 250ul P2 buffer (Lysis buffer)
    • 350ul N3 buffer
    • 750ul PE buffer
    • 500ul PE buffer
    • Columns
  1. Spin cells down at 4000rpm for 10 minutes
  2. Discard supernatant (LB)
  3. Resuspend pellet in P1 buffer
  4. Transfer to labeled Eppendorf tube
  5. Add P2 buffer. Solution should turn blue
  6. Invert tubes 4-6 times, then wait for 2 minutes
  7. Stop the reaction by adding N3 buffer and immediately inverting 4-6 times. Solution should turn clear
  8. Centrifuge at 13000 rpm for 10 minutesv
  9. Decant/pipette supernatant into mini-prep columns. Discard flow-through
  10. Wash with PE buffer (750ul)
  11. Centrifuge at 13000 rpm for 1 minutev
  12. Discard flow-through. Add second wash of PE buffer (500ul)v
  13. Centrifuge at 13000 rpm for 1 minute
  14. Discard flow-through
  15. Centrifuge empty columns at 13000 rpm for 1 minute to eliminate any excess wash buffer
  16. Discard flow-through
  17. Move columns into a labelled eppendorf
  18. Add 30-40ul distilled water and wait for 2-3minutes
  19. Elute DNA by centrifuging at 13000 rpm for 1 minute, do not discard flow-through. Discard column.
  20. Nanodrop


  • Materials
    • 2ul Cutsmart buffer (New England Biolabs)
    • 1ul Restriction enzymes
    • 12-16ul DNA to be digested
    • Distilled water (up to 20ul total volume)
  1. Prepare mix
  2. Incubate for 1-2h at 37 degrees

1% Agarose Gel

  • Materials
    • 1g Agarose
    • 100mL 1X TAE buffer
    • 8uL SYBR Safe
  1. Mix Agarose and 1x TAE buffer
  2. Heat up until Agarose is dissolved
  3. Add SYBR Safe
  4. Pour into gel tray and let cool

Agarose Gel Electrophoresis

  • Materials
    • 1% Agarose gel DNA ladder
    • 6x loading dye
    • Electrophoresis cuvette
  1. Set gel tray into cuvette, filled with 1x TAE buffer
  2. Inoculate samples, previously dyed with 6x loading dye. Additionally, provided a DNA ladder for further reference of DNA sizes
  3. Run gel at 110V for 30-40min

Overnight Cell Incubation

  • Materials
    • 5mL Luria Broth
    • 5ul specific antibiotic
    • Loops (for colony picking or glycerol stock scraping)
  1. Add Luria Broth into 50mL tube
  2. Inoculate specific antibiotic
  3. Scrape/pick glycerol stock surface/colony and transfer into falcon tube
  4. Incubate at 37°C overnight

1x TAE buffer

  • 1x solution contas 40nM Tris, 20mM acetic acid, 1mM EDTA