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Revision as of 17:20, 16 September 2015

iGEM Toulouse 2015

Experiments & Protocols


Content

Sampling of varroa

To run tests with varroas it is necessary to get them back from beehive directly because they cannot live without bees.

Materials

  • Bee hive
  • Beekeeper suit
  • Gloves
  • Smoker
  • Dry twigs
  • Tweezers
  • Big brush
  • Small brush
  • Petri dishes
    Ø x h = 35 x 15 mm

Methods

  1. Slip beekeeper suit and gloves on and go to beehive
  2. Fire dry twigs in smoker
  3. Open bee hive and activate smoker to get bees inside the hive
  4. Take a frame out the hive and remove bees with big brush and smoker
  5. Close beehive
  6. In the lab, put the frame on a table against the wall
  7. With tweezer drill hole into one beehive cell
  8. Remove larvae and look for varroas on larvae and on beehive cell
  9. If there are varroas, take them with a small brush and put them on Petri dishes
  10. Make sure there are two or three larvae on Petri dishes in order to allow survival of varroas
  11. Start again step 7 to 9 until you have enough varroas

Photos 1, 2 & 3: Our teams members gathering varroas on infected larvae

Standardization of varroas and sampling

When we take varroas directly from frame, as it is described in protocol “Sampling Varroas”, we have varroas in different phases. In order to have varroas in the same phase it is necessary to add one step and it is important for reproducibility of the experiments. With this method we place varroas on adult bees so all varroas will be in phoretic phase.

Materials

  • Bees in box with aeration and glucose
  • Varroas from protocol “Sampling varroas”
  • Gas cylinder of CO2
  • Small brush
  • Tweezers
  • Petri dishes
    Ø x h = 35 x 15 mm

Methods

  1. With small brush take varroas from Petri dish and put them on bees in box through aeration holes
  2. Place the box in a 35 °C incubator overnight. Make sure you have a bowl with water in order to have enough humidity in incubator
  3. Take the box out of incubator
  4. Add CO2 from gas cylinder into the box until all bees fall down
  5. Open the box, take a bee with tweezer and look for varroas
  6. When you find a varroa take him with small brush and replace bee in the box
  7. Start again step 5 and 6 until you have enough varroas

Photos 4 & 5: Varroas gathering on infected bees

Attraction test on varroas

In order to test the attraction effect of butyric acid on varroas an Y test was built, as it is showed below. A glass pipe was chosen because on plastic varroas could load themselves with electrostatics and die. For butyric acid, the concentration chosen 4 % (V/V) because this is the concentration used in the patent quoted (see “Attribution” part).

Materials

  • Pump wich expels air
  • 15 mL Flacon tube
  • Plastic pipe
    Ø = 10 mm
  • Glass T pipe
    Ø = 10 mm, made by a glassworker
  • Plastic separator
  • Carded cotton
  • Absorbent cotton
  • 5mL 4% (V/V) Butyric acid
  • 5mL Water
  • Standardized varroas

Methods

  1. Put a cotton on Petri dish and add 400 µL of one acid formic solution
  2. Place three varroas on this Petri dish and close it
  3. Start again step1 and 2 for each formic acid solution and water
  4. Each 30 minutes check if varroas are alive. To do that:
  5. When varroa heads for one side of Glass T tube and covers more than 2 cm test is over and we write down the side choosen by varroa (Butyric acid or Water)
  6. Two tests can be made in the same time thanks to the plastic separator

Photo 6: Glass T-tube

Mortality test on varroas

To test the toxicity of formic acid on varroas, we based our thoughts on present treatments to choose a concentration to use. When beekeepers use formic acid for long treatment they place a diffuser at the top of the hive and formic acid concentration was assessed at 200 ppm 1 on average which is equivalent to 7.8 mmol.m-3. As gas concentration is difficult to evaluate we calculate the liquid concentration balance thanks to the ideal gas law and the Henry’s law. To simplify calculation we noted down formic acid A.

$$ P\cdot V = n\cdot R\cdot T, \textrm{ideal gaz law} $$ $$ P_A = C_A\cdot R\cdot T = 7,826\cdot10^{-3}\times8.314\times293=19,96 Pa $$

  • PA: partial pressure of A in Pa
  • CA: Concentration of A in air in mol.m-3
  • R: perfect gaz constant = 8.314 J.mol-1.K-1
  • T: temperature in °K

$$ P_A = H_A\cdot C_{A,eq}, \textrm{Henry's law} $$ $$ C_{A,eq} = \frac{19,964}{0.019} = 1.019 mol.L^{-1}$$

  • CA,eq: equivalent concentration in liquid in mol.L-1
  • HA: Henry's constant = 0.019 Pa.m3mol-1

So, we chose a positive control with a higher concentration, 2 mol.L-1, and then decreasing concentration in order to identify which minimum concentration could kill varroa. For a negative control we use water. For this test we use varroas from frames directly because we did not have enough standardized varroas.

Materials

  • Petri dishes
    Ø x h = 35 x 15 mm
  • Varroas form “Sampling varroas”
  • Cotton
  • Acid formic solutions:
    • 2 mol.L-1
    • 10 mmol.L-1
    • 1 mmol.L-1
    • 500 µmol.L-1
    • 50 µmol.L-1
  • Water

Methods

  1. Put a cotton on Petri dish and add 400 µL of one acid formic solution
  2. Place three varroas in this Petri dish and close it
  3. Start again step1 and 2 for each formic acid solution and water
  4. Every 30 minutes check if varroas are alive. To do that:
    1. Tap on Petri dish and see if varroa moves. If it does varroa is still alive, if not see below
    2. Observe through a binocular magnifier if varroa move. If it does, it is still alive.

Photo 7: Varroa mortality experiment

Protocols for culture tests

Cytotoxicity tests

Choice of concentrations

In the begining we tested high and low concentrations and in function of results we adapted concentrations. In the end we worked with these concentrations:

  • Butyric acid : 218 mM, 109 mM, 10.9 mM, 5.45 mM and 1.09 mM
  • Formic acid : 100 mM, 10 mM, 1 mM 500 µM, 100 µM, 50 µM and 25 µM

Materials

  • Optical reader, OPTIMA MARS Analysis
  • 48 wells plates
  • Pre-culture of E. coli BW 25113
  • Acid solutions
  • Medium : LB, M9 15 mM of glucose or 30 mM of glucose

Methods

  1. Add 400 µL of medium in each well
  2. Add 50 µL of pre-culture
  3. Add 50 µL of acid solution
  4. Place the 48 well plate in the optical reader
  5. Adjust parameters on computer.
    Usually we set 250 cycles around 24 hours so we have an OD measurement every six minutes

Note: Each condition is tested in three replicates

Culture on erlenmeyers and TubeSpin® Bioreactors

      - Inoculation and sampling

Materials

  • Pre-culture of E.Coli BW 25113 in LB
  • Spectrophotometer
  • 1mL Spectrophotometer cuvettes
  • Centrifuge
  • Erlenmeyers
  • TubeSpin® Bioreactors from TPP brand
  • Medium : M9 15 mM of glucose or 30 mM of glucose
  • Incubators at 37 °C, 130 rpm and without agitation
  • 1.5 mL Eppendorf
  • 0.2 µm filters

Methods

  1. Add 50 mL of medium on erlenmyer and TubeSpin® Bioreactor
  2. Inoculate from pre-culture to have OD600nm=0.1.
    To do that centrifuge the appropriate volume of pre-culture, then remove LB medium and resuspend sediment with M9 medium to inoculate.
    Note: This step permits to eliminate substrates from LB medium which could interfere during NMR analysis.
  3. Place erlenmeyers in incubator 37 °C 130 rpm and TubeSpin® Bioreactor in incubator 37 °C without agitation
  4. Sampling every two hours the first day:
    • Take 1 mL of culture in 1.5 mL Eppendorf.
      For TubeSpin® Bioreactor use needle and syringe in order not to let air enter.
    • Add 100 µL of sample in spectrophotometer cuvette, complete with 900 µL water and measure OD600 nm with spectrophotometer
    • Centrifuge the rest of samples at 13,000 rpm during 3 minutes
    • Filter the supernatant through a 0.2 µm filter and conserve it at -20 °C
  5. Days follow sample once a day with method below and plate on Petri dish an appropriate dilution in order to know if bacteria are alive

      - NMR analysis

Materials

  • Culture supernatants from -20°C
  • 2.5 mM TSP (Trimethylsilyl propanoic acid) diluted in D2O
  • 0.5 mm NMR tubes
  • 1.5 mL Eppendorf
  • Spinners (5mm)
  • 500 MHz Bruker Avance Spectrometer

Methods

  1. Add 400 µL of culture supernatant in 1.5 mL Eppendorf
  2. Add 100 µL of TSP solution
  3. Place the mix in 0.5 mm NMR tubes
  4. Place NMR tube into spinner, sample is ready to analyse

Culture on 48 wells plates

In order to determine the right concentration of polysaccharide and enzyme of BioSilta kit we have to do several cultures at the same time. So, we use an optical reader and 48 wells plates.

Materials

  • Optical reader, OPTIMA MARS Analysis
  • 48 wells plates
  • Pre-culture of E. coli BW 25113
  • Different concentrations of BioSilta medium
  • For one concentration of BioSilta medium different concentrations of enzyme

Methods

  1. Add 450 µL of medium in each well
  2. Add 50 µL of pre-culture
  3. Place the 48 well plate in the optical reader
  4. Adjust parameters on computer. We tested culture between one day and ten days

Note: Each condition is tested almost in two replicates. According to our results we adapt concentrations of Biosilta medium and enzyme, results are exposed in Device part.

Enzyme kinetic

Materials

  • Spectrophotometer
  • BioSilta medium
  • BioSilta enzyme solution named Reagent A (3000U/L)
  • Bradford’s reagent
  • 1.5 mL Eppendorf
  • Standard solutions of glucose

Methods

  1. For each standard solutions : in a 1.5 mL Eppendorf sample 10 µL and add 1 mL Bradford’s Reagent, wait 20 minutes and measure OD505 nm
  2. Plot glucose concentration in function of OD505nm and determine the linear region
  3. Add 1.5 mL of BioSilta medium and 45 µL of reagent A (50 U/L) in an Eppendorf
  4. Sampling every 30 minutes:
    1. Take 10 µL and add 1 mL of Bradford’s reagent in an Eppendorf
    2. Wait 20 minutes
    3. Measure OD505 nm
    4. If OD505 nm is over linear region dilute sample and measure OD505 nm again
  5. Stop sampling when glucose concentration no longer change

Acids production test

In order to test if E.coli produces formic acid and butyric acid with genes added, we made culture test with modified bacteria. We used the same protocol as “Culture on Erlenmeyers and TubeSpin® Bioreactors” with some changes:

  • Volume of culture : 30 mL
  • Add Ampicillin at 25 µg/mL to have selection pressure
  • The number of samples:
    • Sample at the beginning
    • Sample at the end of the first day
    • Sample after 24 hours culture and 48 hours culture

Test of gas concentration

The objective of our device is to produce gas, so we would like to know gas composition of our culture. So, we developed a system in order to recover acids gas.

Materials

  • 4 hours culture in 50 mL Falcon in M9 medium with 15 mM of glucose
  • Silicon plugs adapted to 50 mL Falcon
  • Needles
  • 0.2 µm filters
  • 10 mL Syringes
  • Neoprene pipes Ø=0.8 mm
  • 10 mM NaHCO3
  • 1.5 mL Eppendorf
  • 1 mL Sterile cone
  • Incubator 3 °C without agitation

Methods

  1. Replace Falcon plug with silicon plug
  2. Adjust fliter on needle and peg it into silicon plug. Do it twice
  3. Adjust neoprene pipe into each filter
  4. Add 700 µL of NaHCO3 in an Eppendorf
  5. At the end of first pipe put a sterile cone and immerse it into Eppendorf with NaHCO3
  6. At the end of second pipe put a 10 mL syringe
  7. After 24 hours culture, press 10 mL syringe in order to expel gas in NaHCO3 solution
  8. Conserve samples at -20 °C before NMR analysis (see protocol foregoing)

Note 1: We used culture in M9 because with the “Acids production tests” we had data on this medium.
Note 2: 10 mM NaHCO3 solution was used because pH was 8.3 so it would permit acid gas solubilisation.


Photo 8: Gas concentration test with falcons


Protocol for Polymerase Chain Reaction (PCR), From Thermo Scientific™ DreamTaq™ Green PCR Master Mix

Materials

  • MilliQ water nuclease free (QSP)
  • PCR Mix 2X
  • Forward primer
  • Reverse primer
  • Template DNA
  • Thin walled PCR tube
  • Ice

Methods

  1. Gently vortex and briefly centrifuge the PCR mix after thawing
  2. Place a thin-walled PCR tube on ice and add the different components for a 50 μL PCR reaction
  3. Gently vortex the samples
  4. Perform PCR using the recommended thermal cycling conditions

The PCR Mix from Thermo Scientific contains Taq DNA polymerase, Green Buffer, MgCl2, dNTPs but also two tracking dyes and a density reagent that allows for direct loading of the PCR product on a migration gel.
The template DNA concentration has to be adapted in order to be between 10 pg and 1 μg in the final volume of 50 μL. The template DNA can come from a miniprep solution or from a single colony. The primer concentrations have to be between 0,1 μM and 1 μM.
Each PCR reaction has to be adapted to the length of the PCR products, and to the melting temperature Tm of the primers. The extension step lasts 1 min for PCR products up to 2 kb. For longer products, the extension time has to be prolonged by 1 min/kb.

Step Temperature (°C) Time Number of cycles

Initial denaturation

95

1-3 min

1

Denaturation

95

30s

25-40

Annealing

Tm – 5°C

30s

Extension

72

Adapt to the length

Final extension

72

5-15 min

1

Protocols for TPX® permeability tests

Preparation of TPX® bag

Materials

  • TPX®, gas permeable plastic
  • Fusing machine
  • 2 mM Formic acid solution
  • 4 % (V/V) Butyric acid solution

Methods

  1. Prepare plastic bag in sticking on 3 sides over 4 with fusing machine
  2. Add 7 mL of appropriate solution in plastic bag
  3. Stick on the last side with fusing machine

Permeability test

To test gas permeability of TPX® plastic, we use the same protocol as “Test of gas concentration”. The only change is that no filters were used because the sterility is not necessary.

Photo 9: The device used for the permeability test


Sterility test of TPX® bag (first protocol)

    Demonstrate that the TPX bag is impermeable to bacteria from outside to inside

Materials

  • TPX bags
  • LB Medium
  • E. coli DH5alpha + Psb1c3+rfp
  • Steril laboratory glass bottle

Methods

  1. Overnight culture of E. coli DH5alpha + pSB1C3 + RFP at 37 °C
  2. Fill a small TPX® bag with SOC medium
  3. Put the small bag in a Steril laboratory glass bottle containing an inoculum of DH5alpha in SOC medium + Ampicilline
  4. Negative Control: Fill a small TPX® bag with SOC medium + Amp and put it in a glass bottle which does not contain any bacteria
  5. Incubate at 37 °C
  6. Take a sample and spread in a Petri dish

Sterility test of TPX® bag (second protocol)

    Demonstrate that the TPX® bag is impermeable to bacteria from inside to outside

Materials

  • TPX bags
  • M9 defined Medium
  • E. coli BW 25113
  • Steril laboratory glass bottle

Methods

  1. Overnight culture of E. coli BW 25113 at 37 °C
  2. Inoculate a small TPX® bag at OD600 nm = 0,1 in LB medium (Final Volume = 8 mL)
  3. Negative Control: Fill a small TPX® bag with M9 medium (Final Volume=8 mL)
  4. Dispose each small bag in a Steril glass measuring cylinder containing M9 medium
  5. Incubate at 37 °C
  6. Measure OD600 nm twice a day

Culture test of E. coli in TPX® bag

Materials

  • TPX bags
  • LB Medium
  • Steril clips
  • E. coli BW 25113
  • Steril laboratory flask

Methods

  1. Overnight culture of E. coli BW 25113 at 37 °C
  2. Inoculate a small TPX® bag at OD600 nm = 0,1 in LB medium (Final Volume = 8 mL)
  3. Close the small bag via fusing machine and Put the closed small bag in a Steril laboratory flask
  4. Positive Control: Inoculate a culture tube at OD600 nm = 0,1 in LB medium (Final Volume = 20 mL)
  5. Incubate at 37 °C
  6. Measure OD600 nm twice a day

Transformation Protocol: RbCl method

Media and solution

YETM 500 mL TFB1 200 mL TFB2 200 mL
  • 2.5 g Yeast Extract
  • 10 g Tryptone
  • 5 g MgSO4.7H2O
  • Adjust pH to 7.5 with KOH
  • For Plates: add 7.5 g of Agar
  • 0.59 g KOAc
  • 2.42 g RbCl
  • 0.29 g CaCl2.2H2O
  • 1.98 g MnCl2.4H2O
  • Adjust to pH 5.8 with 0.2 M acetic acid
  • Add dH2O to 200 mL
  • Filter sterilize
  • Store refrigerated at 4°C
  • 0.42 g MOPS
  • 2.21 g CaCl2.2H20
  • 0.24 g RbCl
  • 30 g Glycerol
  • Adjust to pH 6.5 with KOH
  • Add dH2O to 200 mL
  • Filter sterilize
  • Store refrigerated at 4 °C

Preparation of Competent Cells

  • 1. Streak cells froms frozen stock onto YETM plate. Incubate overnight at 37 °C
  • 2. Pick a single fresh colony to inoculate 5 mL of YETM medium. Grow over night at 37 °C.
  • Do not vortex cells at any time after this point in the procedure
  • 3. Dilute 1 mL of culture into 50 mL YETM medium prewarmed to 37 °C
    • Grow at 37 °C for 2 hours with agitation
    • Volumes can be scaled up 5X and all of the 5 mL overnight culture can be used
  • 4. Transfer culture to sterile 50 mL tube. Chill on ice/water 10-15 minutes
  • 5. Centrifuge for 10 minutes at 2,000 rpm at 4 °C. Immediately aspirate off all of the supernatant
  • Do not allow cells to warm above 4 °C at any time in this procedure
  • 6. Resuspend cells in 10 mL of ice-cold TFB1 with gentle re-pipetting. Use chilled glass or plastic pipette
  • 7. Incubate cells on ice for 5 minutes
  • 8. Repeat step 5
  • 9. Resuspend cells in 2 mL of ice-cold TFB2 with gentle re-pipetting. Use micropipet tip (plastic)
  • 10. Incubate cells on ice for 15 minutes
  • Cells may be used for transformation or frozen
    • To freeze: aliquot cell in 200 μL volumes into prechilled 0.5 mL microfuge tube (on ice)
    • Freeze immediately in liquid nitrogen
    • Store cells frozen at -80 °C

Transformation of Competent Cells

  • 1. If starting with frozen competent cells, warm tube/cells by gently twirling between your fingers until just thawed.
    Immedately place on ice for about 5 minutes
  • 2. Add to 1,5 mL eppendorff on ice: 2-3 μL iGEM plate or 1 μL plasmid or 10 μL ligation.
  • 3. Add 100 μL of competent cells and mix by gentle re-pipetting
  • 4. Incubate cells on ice for 20-30 minutes
  • 5. Heat shock the cells exactly 90 seconds at 42 °C
  • 6. Return cells on ice for 2 minutes
  • 7. Add 1 mL of YETM medium. Incubate at 37 °C for 45-60 minutes with slow gentle shaking
  • 8. Plate 0.1-0.2 mL of transformed cells on LB-plate containing the appropriate antibiotic. Incubate overnight at 37°C

Minipreps

  • 1. Resuspend 4 to 12 colonies from the plate and name each colony taken on the tubes and on the plate (A, B, C, …)
  • 2. Resuspend one colony per culture tube in 5 mL of LB medium with antibiotic
  • 3. Let the culture grow overnight at 37 °C in a shaking incubator
  • 4. Use the QIAprep spin Miniprep Kit for each culture tube. The last step consisting in the elution of the DNA is made with elution buffer or water at 55 °C
  • 5. Keep the tubes at -20 °C

Cloning

First step: Digestion

Both parts have the same antibiotic resistance

Vector Insert Digestion control first enzyme Digestion control second enzyme

Volume equivalent to 1 µg of vector miniprep

Volume equivalent to 1 µg of insert miniprep

Volume equivalent to 1 µg of vector miniprep

Volume equivalent to 1 µg of vector miniprep

1 µL of each restriction enzymes

1 µL of each restriction enzymes

1 µL of the first restriction enzyme

1 µL of the second restriction enzyme

2 µL of Fast Digest Green Buffer (Thermo Scientific™)

2 µL of Fast Digest Green Buffer (Thermo Scientific™)

2 µL of Fast Digest Green Buffer (Thermo Scientific™)

2 µL of Fast Digest Green Buffer (Thermo Scientific™)

Up to 20 µL of Milli-Q water

Up to 20 µL of Milli-Q water

Up to 20 µL of Milli-Q water

Up to 20 µL of Milli-Q water

Incubate 15 minutes at 37 °C

The two parts have a different antibiotic resistance

Both parts

Volume equivalent to 1 µg of DNA miniprep

1 µL of each restriction enzymes

2 µL of Fast Digest Green Buffer (Thermo Scientific™)

Up to 20 µL of Milli-Q water

Incubate 15 minutes at 37°C

Migration and gel extraction

  • 1. Prepare a 1 % or 2 % electrophoresis agarose gel with 0.5 X TAE buffer
  • 2. Put 20 µL of sample + 6 µL of marker (1 kb for 1 % gel and 100 pb for 2 %) into the well
  • 3. Migration for 30 min at 100 V or 1 hour at 50 V
  • 4. Bathe 10 minutes in BET
  • 5. Wash in water for 5 minutes
  • 6. The gel extraction is realized thanks to the QIAGEN Gel Extraction Kit

  • Two ways to inactivate the enzymes for the vector
    • Use of DNA Clean up kit for the DNA fragment above 200 pb
    • Heat inactivation at 95 °C for 10 minutes

Second step: Ligation

Mix Negative Control Positive Control

Volume equivalent to 3 molecules of insert (for one molecule of vector)

no insert

Volume equivalent to 3 molecules of insert (for one molecule of vector)

Volume equivalent to 50 ng of digested vector

Volume equivalent to 50 ng of digested vector

Volume equivalent to 50 ng of undigested vector

2 µL of 10X T4 buffer

2 µL of 10X T4 buffer

2 µL of 10X T4 buffer

0.5 µL of T4 ligase

0.5 µL of T4 ligase

0.5 µL of T4 ligase

Up to 20 µL of Milli-Q water

Up to 20 µL of Milli-Q water

Up to 20 µL of Milli-Q water

 Incubate the ligation mix 15 minutes at room temperature (22°C)

 Keep the tubes in ice or at -20 °C to prepare the transformation

Third step: Transformation

  • 1a. Take 10 µL of the ligation mix for 100 µL of competent cells and use the Toulouse iGEM Team 2015 transformation protocol
  • 1b. Positive control: take 10 µL of the ligation mix for 100 µL of competent cells and use the Toulouse iGEM Team 2015 transformation protocol
  • 1c. First restriction enzyme digestion control: take 10 µL of the corresponding digestion mix (First step) for 100 µL of competent cells and use the Toulouse iGEM Team 2015 transformation protocol
  • 1d. Second restriction enzyme digestion control: take 10 µL of the corresponding digestion mix (First step) for 100 µL of competent cells and use the Toulouse iGEM Team 2015 transformation protocol
  • 1e. Negative control: take 10 µL of the corresponding mix for 100 µL of competent cells and use the Toulouse iGEM Team 2015 transformation protocol
  • 2. Plate the solution on selective medium overnight at 37 °C