Team:Alberta/Methodology/Protocols

From 2011.igem.org

METHODOLOGY

Protocols

Growth

Growth in Liquid Culture

  1. Into a 500 mL baffled flask, under a flame, add 100 mL of autoclaved media (PD, VSuTB, or etc)
  2. Add approximately 6 x 106 cells to the each flask.
  3. Incubate the cultures in a rotating incubator at 37 °C and a speed of 200 rpm. Grow until saturation.

Solid Culture Growth

  1. Add 15g/L of Agar to any liquid media recipe, before autoclaving.
  2. Autoclave to sterilize.
  3. While still hot and in liquid form, pour the media into the desired container. 25 mL in a 100 mL conical flask is sufficient.
  4. When the media has polymerized, inoculate the media with a loop of conidia (bright orange in color) from a previous culture or a small amount of conidia suspended in water. Seal the top of the tube with a sponge top.
  5. Incubate the flask at 37 °C for 2 days, then transfer to a well lit room and let the culture conidiate for 5 days at room temperature. It should produce a layer of bright orange conidia, which are ready for harvest.
  6. For growth experiments, harvest this conidia within one or two days to ensure maximum viability. For propagation of a strain, the solid culture can be stored at room temperature for up to 1 month

Harvesting Conidia from Solid Cultures

  1. Add 50 mL of ddH2O to the 100 mL of flask of conidia to be harvested
  2. Swirl the flask a few times to allow for most of the conidia to be suspend. Transfer the suspension of conidia to a 50 mL plastic tube.
  3. Centrifuge the tube at 6000 rpm for 2 min.
  4. Decant the supernatant. Wash the conidia with 40 mL of ddH2O.
  5. Centrifuge the tube again at 6000 rpm for 2 min.
  6. Decent the supernatant. Resuspend the pellet with 10 ml of ddH2O.
  7. Using a hemocytometer and serial dilutions when necessary, count the conidia and estimate the concentration of the tube.
  8. Dilute the conidia to the desired concentration.

Potato Dextrose Media (PD)

  1. Take 300 g of Potato (Baking Potatoes) and dice into 0.5 cm cubes.
  2. Add 1.0 L of water and boil for 30 min.
  3. Strain out the potato chunks and add 20 g of dextrose (D-glucose)
  4. Autoclave solution
  5. For solid PD agar, add 15 g of agar per 1 L of media.

VSuTB (Vogel's Sucrose Trace Elements and Biotin) Media

The following protocol was given to us from the Nargang Lab at the University of Alberta. It is based off the protocol listed on www.fgsc.net/methods/vogels.html.


VSuTB 50 mL 100 mL 500 mL 1000 mL
50x Vogel's Salts 1 mL 2 mL 10 mL 20 mL
15g/L Table Sugar 0.75 g 1.5 g 7.5 g 15 g
100mg/L Trace Elements 50 µL 100 µL 0.5 mL 1 mL
100mg/L Biotin 50 µL 100 µL 0.5 mL 1 mL
15g/L Agar 0.75 g 1.5 g 7.5 g 15 g
dH2O 48.9 mL 97.8 mL 489 mL 978 mL
  • For liquid VSuTB media, exclude the agar.
  • Mix all the ingredients in a proper container. Autoclave to sterilize.
  • The composition of the 50x Vogel's Salts, Trace Elements and Biotin solutions are listed on www.fgsc.net/methods/vogels.html. The above protocol deviates for 50x Vogel's Salts; the Trace Element and Biotin solutions are added separately as individual components to make the final Vogel's media.

Coffee Grounds Media (10% Wet Weight)

  1. Take 50 g of Coffee Ground from Starbucks
  2. Add 500 mL of dH2O and autoclave to sterilize

Sawdust Media (5% dry weight)

  1. Take 25 g of sawdust (source: RONA woodshop)
  2. Add 500 mL of dH2O and autoclave to sterilize

Fertilizer solution

  • Prepared 10X concentrated as per the dilution recommend by the supplier.

1st Wheat Straw Media

  1. 10 g wheat straw cut into ~2 cm pieces
  2. Add 500 mL of dH2O and boiled for 30 mins
  3. Strained 250 mL with a kitchen strainer and left the other 250 mL unstrained.
  4. Autoclaved both to sterilize.

1st Grass Media

  • Same as 1st wheat straw media, except grass was dried on a hot plate first (10 g grass)

Wheat Straw with 1% w/v NaOH treatment

  1. 1 g of ground wheat straw (using food processor until it passed through the pores of a kitchen strainer)
  2. Add 100 mL of 1% NaOH, autoclaved
  3. Washed the wheat grounds on filter paper with dH2O until pH 7.0.
  4. Add 100 mL of dH2O to the neutralized grounds and autoclaved to sterilize.

10X MgCa solution

  1. Dissolve 16.6 g of MgCl2* 6 H2O and 10.0 g of CaCl2 * 2 H2O in 100 mL of dH2O.
  2. Mix under totally dissolved
  3. Autoclave to sterilize.

Baked Sawdust

  1. Baked sawdust in an oven at 400oF for 1 hr.
  2. Take 2 g of the baked sawdust, and add 200 mL of dH2O.
  3. Boil for 30 min. Strained out the sawdust for half the solution; leave the other half in a separate bottle.
  4. Autoclaved both bottles.

Race Tube

  • Race tubes were used to show amount of growth by hypheal extension.
  • Race tubes were custom-made by the University of Alberta Glass Shop.
  • Our race tubes were 1 meter in length with an interior diameter of 25 mm.

  1. For each of media, stopper one tube with a rubber stopper.
  2. After autoclaving, while it is still hot (and liquid), pour the media in the tube. And stopper the other end with a rubber stopper.
  3. Allow the media inside the tube to cool and the agar to polymerize.
  4. When the agar is solid, remove a rubber stopper at ONE end of the tube and immediately replace with a sponge stopper.

WS + FBT 3 g wheat straw, 100 µL trace elements, 100 µL biotin, 10 mL MgCa solution 10 mL fertilizer, 1.5 g agar, 80 mL dH2O.
WS + F 3 g wheat straw, 10 mL MgCa solution, 10 mL fertilizer, 1.5 g agar, 80 mL dH2O.
GS + FBT 3 g grass clippings, 100 µL trace elements, 100 µL biotin, 10 mL MgCa solution 10 mL fertilizer, 1.5 g agar, 80 mL d H2O.
GS + F 3 g grass clippings, 10 mL MgCa solution, 10 mL fertilizer, 1.5 g agar, 80 mL dH2O.
Vogel's media prepared as listed above and add a few drops of food colouring to add contrast from Neurospora crassa in race tube experiments

Genetics

PCR Clean-up Protocol

  1. Add 5X the volume of buffer PB, and ~10µL of 3M sodium acetate and mix by inverting.
  2. Apply the mixture to filter-columns that have been placed on the vacuum manifold.
  3. Wash the columns by adding 750 uL Buffer PE and applying vacuum. Wait until all liquid is gone and then turn vacuum off.
  4. Place each column in the 2 mL rounded collection tubes and centrifuge at 13000 rpm for 1 min
  5. Discard the 2 mL collection tubes and place your columns into labelled 1.5 mL eppendorf tubes.
  6. Apply 30 uL of Sterile TE Buffer (10 mM EDTA) to the CENTER of each column
  7. Let stand for AT LEAST 10 MINUTES
  8. Centrifuge for 1 min at 13000 rpm.
  9. Discard the column into the discarded column beaker.

Miniprep Protocol

  1. Pellet your overnight culture tubes by adding ~1800 uL of each culture to separate 2mL eppendorf tubes, centrifuging at 13000 rpm for 7 minutes, discarding the supernatant (just pour it out), and adding more culture. Centrifuge again, discard the supernatant, and if there’s any culture left, add it and centrifuge again. Discard the final supernatant.
  2. Resuspend each pellet in 250 uL of Buffer P1. Vortex to ensure complete resuspension
  3. Add 250 uL Buffer P2, and immediately mix thoroughly by inverting ~7 times. Do NOT leave this mixture stand for more than 5 minutes. Preferably, move on to step 3 within 2 minutes or so.
  4. Quickly add 350 uL Buffer N3. Mix a few times via inversion.
  5. Centrifuge all tubes for 10 min at 13000 rpm
  6. Apply the supernatant from each tube to 4 LABELLED filter-columns that have been placed on the vacuum manifold. Once all samples have been applied to the column, apply the vacuum to the manifold. Let the samples drain through until no liquid is left.
  7. Wash the columns by adding 750 uL Buffer PE and applying vacuum. Wait until all liquid is gone and then turn vacuum off.
  8. Place each column in the 2 mL rounded collection tubes and centrifuge at 13000 rpm for 1 min
  9. Discard the 2 mL collection tubes and place your columns into labelled 1.5 mL eppendorf tubes.
  10. Apply 50 uL of Sterile TE Buffer (10 mM EDTA) to the CENTER of each column
  11. Let stand for AT LEAST 10 MINUTES
  12. Centrifuge for 1 min at 13000 rpm.
  13. Discard the column into the discarded column beaker.

Creating Parts Plasmids

  1. Ligate together the correct prefix, (suffix, if needed), cut part, and cut plasmid backbone.
    • 2 uL of cut 0.025 pmol/uL part
    • 2 uL of cut 0.025 pmol/uL plasmid
    • 2 uL of 0.025 pmol/uL prefix
    • (2 uL of 0.025 pmol/uL suffix)
    • 2 uL of 5X T4 DNA Ligase Buffer (in PEG)
    • 1 uL T4 DNA Ligase.
    • Add Mq’d H2O to 10 uL (or just add none)
    Ligate anywhere between an hour and overnight
  2. Transform the entire 10 uL of ligation reaction into 100 uL of competent cells per the usual transformation protocol. Plate on chlor-inoculated plates.
    • Chlor plates have 4.0 mL chloramphenicol/L of LB agar
  3. The next day, pick 4 individual white colonies and inoculate into sterile culture tubes with 4 mL LB medium and chlor at 4 mL/L. Shake at 37oC for ~16 hours (overnight).
  4. Miniprep the overnights to get 4 concentrated tubes of YOUR particular parts plasmid.
  5. Check the concentration of your minipreps. If they look ok (i.e. high), then go ahead and digest some of MP#1 with NotI, and some of MP#1 with BsaI.
    • NotI Digestion protocol:
      1. Add 1 ug of your DNA to a 1.5 mL eppendorf tube (will be a variable volume)
      2. Add 5 uL of 10X NEBuffer 3
      3. Add 5 uL of 10X BSA
      4. Add 1 uL of NotI
      5. Add Mq’d H2O to a total volume of 50 uL (be sure to mix)
      6. Incubate at 37°C for at least 1 hour. Can incubate overnight.
      7. PCR clean-up, and elute in 30 uL TE.
    • BsaI Digestion Protocol:
      1. Add 1 ug of your DNA to a 1.5 mL eppendorf tube
      2. Add 5 uL of 10X NEBuffer 4
      3. Add 5uL of 10X BSA
      4. Add 1 uL of BsaI-HF
      5. Add Mq’d H2O to a total volume of 50 uL.
      6. Incubate at 37°C overnight.
      7. PCR clean-up, and elute in 30 uL TE
  6. Run on a gel the following samples:
    • Lane 2: The pure Miniprep that you took some of to digest with (MP #1)
    • Lane 3: MW Ladder
    • Lane 4: The results of your NotI digestion
    • Lane 5: The results of your BsaI digestion

Extraction and Esterification

Sample Preparation for GC Analysis:

  1. wash clean glass tubes with a 2:1 chloroform:methanol solution; drain solution and let evaporate in fume hood
  2. set water bath to 70 degrees Celsius
  3. weigh out day 5 N. crassa samples (~0.1-0.3g)
  4. transfer samples to clean class tubes; add 2 mL of methanolic HCl to each sample
  5. incubate samples in glass tubes in a 70 degree water bath for one hour
  6. add 0.9% NaCl to each rxn tube and mix (rxn is stopped)
  7. add 2 mL hexane_IS (0.5 mg/ml C15) to each glass tube, vortex 2 min @ max speed
  8. spin @ 3000 rpm for 5 mins
  9. carefully take the tubes out and transfer the top phase (hexane_IS) into a fresh glass tube; IMPORTANT: do not disturb the intermediate layer

(Hexane_IS NU-CHEK Prep Inc. LOT# A-615-J30-)

Production of raw fuel:

  1. wash clean glass tubes with a 2:1 chloroform:methanol solution; drain solution and let evaporate in fume hood
  2. set water bath to 70 degrees Celsius
  3. weigh out day 5 N. crassa samples (~10-30g)
  4. dried out large mass of N. crassa and crushed with a motor and pestle
  5. add the N. crassa to a large screw top jar, and add methanol and methanolic HCL
  6. use 10x ml of liquid for mass of fungus. ¾ of liquid volume should is methanol, ¼ is 3M methanolic HCL.
  7. incubate samples in glass tubes in a 70 degree water bath for two hours
  8. add ½ the volume of the reaction of dd H2O
  9. add ½ the volume of hexane, and shake vigorously
  10. Let settle and pipette off the top hexane layer, being careful not to take any of the bottom layer.
  11. repeat steps ix-x, and add to the rest of the hexane
  12. remove hexane under negative pressure.

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