Team:USP SaoCarlos-Brazil/Experiments

IARA

EXPERIMENTS

OVERVIEW

The experiments revolves around the assembly and construction of the following biosynthetic circuits, divided in three generations:

DESCRIPTION

The first experimental steps consists in the construction of the main circuits Secretion System and MermAID in iGEM vector pSB1K3, along with the di-guanylate cyclase assemblies in the plasmid pETSUMO. They constitute our first generation, and the others assemblies are derived mainly from the MermAID and a choice of the best di-guanylate cyclase.

Vectors mermAID
Secretion System

The second generation relies in the amplification of our composite part MermAID and only the chimeric protein Iara-alpha and it’s insertion in the pACYCDuet vector, which is regulated by lac promoter and allows a future co-transformation with the Secretion System. The planned grand finale was the insertion of the di-guanylate cyclase that stood out during tests along with Iara-alpha in pACYCDuet. This plasmid then would be ready to be co-transformed with Secretion System, forming the complete biosynthetic system.

To prove that the protein Iara-α is in fact capturing and binding to mercury, we proposed two experiments that would give us an indirect answer. Our work hypothesis is: if bacterias contain the synthetic biological circuit and are expressing it, a biological advantage would be observed through it’s resistance and consequently would happen colony growth in a medium containing a concentration of mercury. The first experiment was made according to the protocol Hg Disc Diffusion Test and the second consists in a growth curve, following the protocol Growth curve & MIC determination. The biofilm formation was verified through tests that induced it’s production and subsequently stain by violet crystal.

Synthetic DNA ordered from TwistBioscience

OBS: Our project begun three months after what was planned due to delay in the arrival of our synthetic DNA. Besides the time it took to arrive at Brazil, our DNA was retained by ANVISA, Brazil’s National Health Surveillance Agency.

LB Medium and LB Medium Solid

Our LB Medium is from Sigma-Aldrich (LB Broth - Sigma L3022-1KG), just dilute it in distilled water. Concentration: 20g/L

Materials:

  • Beaker
  • Graduated beaker
  • Magnetic stir bar
  • Distilled water
  • LB medium
  • Agar * (solid medium)

  1. Weigh half LB in a beaker (small volume);
    For solid medium for plates, add 2% (w / v) agar;
  2. Add less volume than total distilled water;
  3. Place on the shaker with a magnetic stir bar until smooth;
  4. Transfer to a graduate beaker;
  5. Make up to volume with distilled water;
  6. Return to beaker and mix on shaker until smooth again;
  7. After preparation, autoclaving the medium.

Preparation of Antibiotic Stock Solutions

REAGENT

DILUTION IN

[]STOCK*

[]USE
Kanamycin Water 100 mg/ml 50 ug/ml
Chloramphenicol Ethanol 68 mg/ml 34 ug/ml
*1ml of 2000x concentrated stocks (eppendorf)
NOTE: Concentration indicated on eppendorf as ‘x’ mg (‘x 'mg / ml).
  • Chloramphenicol (C28)
    NOTE: Chloramphenicol a light sensitive. Avoid exposing it to light!
    1. Weigh mass to desired volume of solution;
      1. Beaker
      2. Falcon
    2. Add volume, less than total of ethanol (95%);
    3. Dissolve completely;
      1. Magnetic stir bar
      2. Vortex
    4. Transfer to a graduated beaker;
    5. Make up to volume with ethanol (95%);
    6. Distribute 1ml for each eppendorf stock;
    7. Cover with aluminum foil;
    8. Store at -20ºC.
  • Kanamycin (K1)
    NOTE: Poorly soluble. Attention to homogenize.
    1. Weigh mass to desired volume of solution;
      1. Beaker
      2. Falcon
    2. Add volume, less than total of sterile water;
    3. Dissolve completely;
      1. Magnetic stir bar
      2. Vortex
    4. Transfer to a graduated beaker;
    5. Make up to volume with sterile water;
      (In the flow):
    6. Pass solution through the syringe filter (eliminate possible microorganisms);
    7. Distribute 1ml for each eppendorf stock;
    8. Store at -20ºC.

Preparation of Plates with Medium and Antibiotic

Comments:

  • Always work with autoclaved medium;
  • Only open autoclaved medium inside the flow;
  • The addition of antibiotic to make the medium selective is done only at the time of use the medium (antibiotic cannot be autoclaved, because it degrades him).
  • Agar-containing medium (solid medium), after liquefying, rapidly gets hard at room temperature. Fast handling required.
Preparation in the flow:
  1. Melt solid medium (LB + agar, autoclaved).
    1. Semi-screw cap: unscrew 1/2 turn;
    2. Microwave for 1 minute;
    3. Shake to help the liquefaction;
    4. If necessary, place for another 20' in the microwave.
      ** Only glass containers go inside the microwave (falcon tube CAN NOT go in the microwave).
      ** Keep an eye for the medium not to boil and overflow. If it begins to bubble very close to the lid, stop the microwave cycle.
  2. Add antibiotic (make selective medium).
    1. Work Concentrations:
      1. Kanamycin: 50ug / ml
      2. Chloramphenicol: 35ug / ml (light sensitive: cover with Al paper)
    2. Check the total volume of medium to be worked with, to get the correct volume of antibiotic solution;
    3. Place medium (liquid), in falcons tubes, 1 for each antibiotic to add them in the medium;
    4. Wait for the moment when the tube containing the liquid medium can be held directly by hand, to add the antibiotic (temperature at which the antibiotic will not be degraded).
  3. Plate the medium in petri plates.
    ** Minimum of medium per plate: 15ml.

Pre-inoculum preparation

NOTE: If the pre-inoculum is not done on the day immediately after plating, the plates with the colonies will be stored in the refrigerator at 4 ° C.

Materials:

  • Autoclaved test-tubes;
  • Liquid medium (selective, in case of transformed cells)
  • Transfer handle

(Preparation in the flow)

(use the Bunsen burner)

(Always flambé the outlet of the glassware/handle).

  1. Addition of culture medium to the test-tube (10 ml);
  2. Flambé handle (wait for it to cool within the safety zone of the Bunsen burner);
  3. Collect 1 transformed colony;
  4. Insert the handle into the test-tube containing medium (ensure that the liquid medium has touched the point of the handle where the colony was);
  5. Thread the test tube partially for aeration (½ turn);
  6. Overnight growth (16h), shaking at 37°C (recommended masking tape to hold the tube cap).

DNA resuspension iGEM kit

Every plate in the kit comes with a aluminium foil protection in order to avoid cross contamination between samples. Never remove this protection. Always manipulate plates in the flow chamber:

  1. Using a pipette tip, make a small hole on the well’s lid;
  2. Add 10 uL of distilled water;
  3. Carefully resuspend the DNA with the pipette;
  4. Let the DNA rest for 5’ at room temperature for hydration;
  5. DNA is ready to be used!
  6. After the resuspension, store the plate at -20oC
  7. Use 1uL to perform a transformation.

Bacterial transformation 1

Manipulation in the flux and on ice:

  1. Add 100 uL of competent cell in an autoclaved eppendorf (1.5 mL);
  2. Add DNA (50 - 100 ng);
  3. Ice 30’;
  4. Thermal-shock 42ºC, 1.5’;
  5. Ice 5’;
  6. Add 250 uL sterile LB medium (without antibiotic);
  7. Incubate for 45’, 37ºC, under agitation (150 rpm);
  8. Plaque transformed cells in selective medium;
  9. Incubate overnight, 37°C, static;

Miniprep - Plasmid DNA Extraction (CellCo)

NOTE: CellCo Fast-n-Easy Plasmid Mini-Prep Kit. The kit comes with all the necessary solutions/buffers, plus the column and collection tubes (fit into the column).

(Cell Lysis)

  1. Add aliquot of cell culture (1 - 3 ml) in an eppendorf;
  2. Centrifuge;
  3. Add 300uL of 'Lysis Buffer';
  4. Resuspend in the pipette for 1'.
    1. Neutralization
  5. Add 300uL of 'Neutralization Buffer';
  6. Mix gently by inversion, maximum of 4-6 times;
  7. Centrifuge in the 10,000g microcentrifuge for 5'.
    1. Yellow color: correct pH, proceed
    2. Purple, orange color: adjust pH.

(Column Activation)

  1. Add 100uL of 'Activation Buffer' to the column;
  2. Place column in a 2ml collection tube;
  3. Centrifuge in the 10,000g microcentrifuge for 30''.

(Column loading)

  1. Add supernatant (7.), to the column;
  2. Centrifuge in the 10,000g microcentrifuge for 30'';
  3. Discard the eluate;
  4. Place column in a 2ml collection tube;

(Column wash)

  1. Add 500uL of 'Wash Buffer';
  2. Centrifuge in the 10,000g microcentrifuge for 30'';
  3. Discard the eluate;

    * For super purification, extra steps:

    1. Add 700uL of 'Wash Buffer';
    2. Centrifuge in the 10,000g microcentrifuge for 30'';
    3. Discard the eluate;
    4. Centrifuge in microcentrifuge 10,000g for 2';

(Elution)

  1. Place column in a clean 1.5ml microtube;
  2. Add 50uL of Milli-Q water;
  3. Inoculate at room temperature for 1';
  4. Centrifuge in the 10,000g microcentrifuge for 1' to elute the DNA.

Oligos resuspension (EXXTEND)

Preparation of Resuspension Solution

First, prepare Tris 1M and EDTA 0.5M, according to the following protocols:

  • 1M TRIS (Tris (hydroxymethyl) aminomethane) (pH = 8.0) (Volume = 1L):
    • Base Tris: 121.12g;
    • Adjust pH 8.0 with HCl;
    • Complete the sufficient amount for 1000 ml with Milli-Q H2O.
  • EDTA (Ethylenediaminetetraacetic Acid) 0.5M (pH = 8.0) (Volume = 50ml):
    • EDTA: 9.305g (MW = 372.22);
    • Adjust pH = 8.0 with concentrated NaOH;
    • Complete the sufficient amount for 50 ml with Milli-Q H2O.

Autoclave EDTA and Tris solutions before preparing TE 1X buffer!

  • 1X TE Buffer Solution [10 mM Tris-HCl, 0.1 mM EDTA (pH 7.5 - 8.0)] (Volume = 500ml)
    • Add 5 ml of 1M Tris;
    • Add 0.1 ml of 0.5M EDTA;
    • Add 494.9 ml of Milli-Q H2O.

Store 1X TE at room temperature in a closed bottle with a lid and parafilm.

Resuspension of the oligos

Exxtend synthesized oligos are shipped dry as they are highly stable in this form.

(After receiving the oligo):

  1. Centrifuge the tube to make sure that any dry oligo that has come off the tube bottom during shipment is returned to it;
  2. To resuspend the oligo use TE buffer solution (10 mM Tris, 0.1 mM EDTA, pH 8.0);
  3. Let the oligo hydrate for a few minutes with intermittent vortex before quantification.

** It is important to note that dilution of the stock solution to working concentration should always be done in pure DNA free water.

Preparation of 100 µM oligo stock solution

Exxtend provides an oligo specification sheet stating quantitative data in nmoles, optical density (OD) units, micrograms (µg) and molecular weight (PM). This allows you to calculate the stock solution at any concentration value you prefer. The general rule is that for any oligo, the number of nmoles x 10, will give you the amount of solvent, in microliters, to obtain 100 µM solution (pmol / µL).

[Molar Concentration] x [Volume] = [mol number], where [1 nmol = 1000 pmol]

Therefore,

[Volume in µL] = [nmol] x [1000 pmol / 100 µM (pmol / µL)] = nmol x 10

The oligo specification sheet also shows the dilution volume in µL for 100 pmol/µL (µM) concentration. Using this volume of solution, your oligo will be suspended at the stock concentration (100 pmol/µL = 100 µM).

Remember that the oligos should be diluted in TE solution (10 mM Tris, 0.1 mM EDTA, pH 8.0), and diluted to working concentration (5-10 µM) in pure water free of DNAses.

Synthetic DNA Resuspension (Twist Bioscience)

NOTE: Twist Bioscience products are dried and shipped in 2ml microcentrifuge tubes or 96-well plates. Although double stranded DNA and single stranded oligonucleotides are stable under most standard laboratory storage conditions, it is important to consider the following best practices to maintain the high quality of Twist Bioscience synthesized DNA.

  1. Upon receipt, centrifuge the tube or plate briefly and resuspend in nuclease-free Tris-EDTA (TE) buffer, pH 8.0 or 10 mM Tris-HCl, pH 8.0 to the desired concentration. We do not recommend resuspension in water.
  2. A concentration of at least 10 ng/uL is recommended for stock dilution, but the optimal concentration will need to be determined based on the application of interest.
  3. Prepare aliquots of stock dilution solution and separate working aliquots to limit the chance of contamination and reduce the number of freeze/thaw cycles. Use working aliquots as soon as possible after preparation and minimize exposure to high temperatures.
  4. For long term storage (over 1 year), freeze the DNA at -20 °C. For even longer storage, freeze the DNA at -80 °C.

Preparation 0.8% agarose gel

NOTE: Due to the use of ethidium bromide in the agarose gel, the entire gel preparation step should be performed in the BrEt specific laboratory area. Always handle nitrile gloves. Do not leave the specific area with the gloves employed in the process.

(In the BrEt area):

  1. Prepare the gel base on the stand by leveling it;
  2. Choose the comb with the desired amount/size of wells and fit into the base of the gel;
  3. Weigh agarose in the conical flask;
  4. Add 1X TAE buffer volume;
    1. If missing, dilute TAE 25X Buffer: 24 distilled water to 1 TAE 25X;
  5. Solubilize agarose, use of microwaves;
    1. 30 seconds in the microwave;
    2. Shake erlenmeyer;

      * Hot glassware, handle with the help of a paper towel;

      1. If visible agarose particulate matter, repeat microwave oven every 10 seconds, being careful not to boil the mixture;
      2. Cool the mixture under tap water by shaking the conical flask until it can be held by hand;

        * Do not overcool to prevent early solidification.

        1. Add ethidium bromide;
        2. Stir to homogenize;
        3. Pour the mixture into the base of the gel;
          1. If there are bubbles, with a tip, take them to the end of the gel removing them from the race path.
        4. Wait for gel to harden until opaque (10~15 minutes);
        5. Put the base in the tub

          * Place the gel with the wells side to the same side of the black electrode.

          1. Fill the bowl with TAE 1X to the maximum height mark;
            1. Ensure that the gel is completely covered.
          2. Connect the cover wires to the source (black to black and red to red);
          3. Turn on the source (switch);
          4. Set the desired parameters;
          5. Put the lid on the bowl;
          6. Press the ON/OFF button to start the race;
            1. If all is correct, small bubbles will rise on both sides of the tub.
          7. After the race is over, press the ON/OFF button to stop and remove the cover.

Purification of PCR product from agarose gel

NOTE: Promega Kit 'Wizard SV Gel and PCR Clean-Up System'. The kit comes with all the necessary solutions, besides the column and collection tube (fits the column).

NOTE 2: Due to the use of ethidium bromide in the agarose gel, the entire gel manipulation step should be performed in the BrEt specific laboratory area.

(Agarose Gel Dissolution)

(Specific Area BrEt)

  1. Extract gel band with corresponding DNA (cut with blade in UV transilluminator);

    * Minimum possible UV time (avoid sample degradation).

    1. Weigh gel slice:
      1. Tare the eppendorf of 1.5 mL;
      2. Add gel slice to eppendorf;
      3. Obtaining the extracted gel mass.
    2. Add 10 μL of Membrane Binding Solution to each 10 mg of gel;
    3. Vortexing;
    4. Incubate at 50-60 °C until the gel has completely dissolved;

(DNA binding to column)

  1. Place a mini column in a collection tube;
  2. Add all volume of gel to the column and incubate at room temperature for 1 minute;
  3. Centrifuge at 16000 g or 14000 rpm for 1 minute;
  4. Remove column from collection tube and discard passing liquid;
  5. Return the column to the collection tube;

(Column wash)

  1. Add 700 µL of 'Membrane Wash Solution';
  2. Centrifuge at 16000 g or 14000 rpm for 1 minute;
  3. Discard the passed liquid;
  4. Add 500 µL of 'Membrane Wash Solution';
  5. Centrifuge at 16000 g or 14000 rpm for 5 minutes;
  6. Discard the passed liquid;
  7. Centrifuge with the centrifuge lid open for 1 minute, for ethanol evaporation (IMPORTANT);

(DNA Elution)

  1. Transfer the mini column to a clean 1.5 mL eppendorf;
  2. Place 50 µL of free-water nuclease (heated to 60 °C), directly in the center of the column but not touching the membrane.

    * Do not use the nuclease free-water kit.

    1. Incubate at room temperature for 1 minute;
    2. Centrifuge for 1 minute at 14000 rpm;
    3. Discard mini column and store DNA at 4 °C or -20 °C.

Calcium chloride competent cells

CaCl2 Solution - pH 7.0

CaCl2 60mM (0,882g if di-H2O)
HEPES 10mM (0,238g)
Glycerol (100%) 15% (15 ml)
H2O Complete to 100 ml

(Day 1 - Preparations):

  1. Streaking in LB the cell of interest (DH5ɑ / BL21);
  2. Autoclaving 100 ml of liquid LB;
  3. Autoclaving 100 ml CaCl 2;
  4. Autoclaving 2 centrifuge tubes of 250 ml;

(Day 2 - Preparations):

  1. Make a 5 ml post-inoculum with a plate colony and leave overnight at 37 °C and 250 rpm;

(Day 3 - Competent Cells):

(On ice! Work with everything cold, including the solution)

  1. Place 2 ml pre-inoculum in the 100 ml medium and incubate at 37 °C and 250 rpm until OD = 0,375;
  2. Distribute the culture in 2 centrifuge tubes (cold), and centrifuge at 1600g for 7min and at 4ºC;
  3. Resuspend the pellet in 10 ml of CaCl2 solution, shake with circular movements in the hand (cells are fragile);
  4. Resuspend the pellet in 10 ml of CaCl2 solution and incubate for 30 min on ice;
  5. Centrifuge again at 1100g for 5min and at 4 °C;
  6. Resuspend the precipitate in 1 ml of cold CaCl2 solution;
  7. Transfer to a sterile 1.5 ml eppendorf;
  8. Aliquot 50ul;
  9. Freeze in liquid nitrogen;
  10. Store at -80ºC.

PCR - Q5 High-Fidelity 2X Master Mix (NEB)

The protocol used for this PCR can be found in the official NEB website (https://international.neb.com/), and it is this one:

https://www.neb.com/protocols/2012/12/07/protocol-for-q5-high-fidelity-2x-master-mix-m0492

Biofilm quantification using violet crystal 2.0

A three days protocol.

  • Day 1: Preparations
    1. Pre inoculum (5mL LB medium + antibiotic), overnight growth, 37ºC, under agitation;
    2. Medium SOC liquid, including erlenmeyers (125 mL) with 20 mL for each construction;
    3. Violet crystal 0,1% solution;
    4. Acetic acid 30% solution;
    5. PBS buffer;
    6. Autoclaved water;
    7. Sterile 24-well plate;
    8. Sterile (or not) 96-well flat bottom plate.
  • Day 2: Experiment set up
    1. Pre inoculum’s OD measurement (600nm)
      1. Dilution 1:1 in LB medium for measurement in spectrophotometer
    2. Calculations for culture’s inicial OD (60nm) = 0.1, in the final volume of 20 mL of SOC medium (erlenmeyer 125 mL):
      ODincitial ✕ V mL = 0.1 ✕ 20 mL
      V: pre inoculum’s volume
    3. Inoculation of SOC medium prepared in erlenmeyer (125 mL), shaker 150 rpm, 37ºC, about 4 hours for growth rate conference;
      Biofilm production
    4. Adding cells and SOC medium + antibiotic in the plate, so that the culture starts it’s biofilm production with 2 mL total volume, OD (600nm) = 0.2;
      1. Calculation of the inoculum volume:
        ODinitial ✕ V mL = 0.2 ✕ 2 mL
        V: inoculum’s volume
      2. Complete to 2 mL with SOC medium + atb
    5. Example of 24-well plate assembly:
      Blank Control Control Control Control Blank
      Blank Samples A Samples A Samples A Samples A Blank
      Blank Samples B Samples B Samples B Samples B Blank
      Blank Samples C Samples C Samples C Samples C Blank
      1. Blank: 2 mL SOC + atb
      2. Control: Culture without the plasmid in SOC medium + atb
      3. Samples: V(DO600 = 0.2) Culture + V (to 2 mL) SOC medium + atb
        ** In case of lac operator, add 2 uL IPTG (1M)
    6. lncubate for 24 hours, 37ºC, under agitation (150 - 180 rpm);
    7. After 24 hours, begin the biofilm quantification.
  • Day 3: Biofilm quantification
    1. Pour the plate in a discard reservatory with sodium hypochlorite;
    2. Wash gently with PBS 3 times;
    3. Let it dry at room temperature. The excess can be removed with absorbent paper and then let the plate dry face up;
    4. Add 2 mL methanol (in fume hood), close the plate and 15 minutes in refrigerator (~15ºC);
    5. Pour the methanol (in fume hood) in an proper discard;
    6. Press the plate against an absorbent paper and let it dry face up to evaporation of the fixer;
    7. Add 2 mL violet crystal, wait 20 minutes;
    8. Pour the violet crystal and use water to wash gently the plate;
    9. Remove water excess with an absorbent paper;
    10. Add 2 mL acetic acid 30% and homogenize each well;
    11. Transfer 125 uL of each well to a 96-well flat bottom plate for Abs reading in 590nm.

Preparation mercury (Hg) solutions

Attention: mercury is toxic. Working with it requires safety measurements: EPI’s.
Solutions of 1ml in 3 concentrations: 20 mg/mL, 200 ug/uL and 20ug/uL of the mercury salt mercury chloride (HgCl2).

  • Solution [HgCl2] = 20 mg/mL
    1. Weight the mass of HgCl2: 20 mg;
    2. Add 1 mL water (Milli-Q);
  • Solution [HgCl2] = 200 ug/mL
    1. Dilute the 20 mg/mL solution: in a eppendorf (1.5 mL) add 10 uL of 20 mg/uL solution and complete with 990 uL of water (Milli-Q);
  • Solution [HgCl2] = 20 ug/mL
    1. Dilute de 200 ug/mL solution: in a eppendorf (1.5 mL) add 100 uL of 200 ug/uL solution and complete with 900 uL of water (Milli-Q);

Hg Disk Diffusion Test

Obs: protocol developed by the team in order to confirm that transformed bacteria are indeed able to survive in Hg contaminated media.

Materials:

  • paper filter cut in circles of ~10mm diameter
  • Hg salt (HgCl2)
  • MilliQ water
  • Pre-inoculated transformed bacteria
  • Petri dishes

Methods:

  1. Set pipette for 10 uL to soak paper filter disc in Hg solution
  2. While the disc dries, spread pre-inoculated cells over prepared Petri dish
  3. Using a sterilized tweezers, pinch Hg dripped disc
  4. Carefully place it over the agar medium
  5. Close plate and let it incubating for the experiment time span of interest.

Quantification of coconut fiber biofilm

Materials:

  • PBS;
  • methanol;
  • 30% acetic acid;
  • Violet crystal 0.1%;
  • Plate with 24 wells (fiber);
  • Flat plate with 96 bottom wells.

(Qualitative visual analysis: photos):

(Quantification):

  1. Fiber-board separation:
    1. Remove the fiber from each well and place it in a new plate with 24 wells;
    2. Pour the plate into an ice cream pot (with hypochlorite - bleach).
  2. Wash gently with 3X PBS buffer;
    1. ** For fibers, remove one by one and position in the respective location of the well on the lid. Wash the plate and then the fiber one by one, returning them to their respective wells.
  3. Allow to dry at room temperature;
  4. Add 2ml methanol (chapel), (total 32ml / plate). Put the lid and than in the refrigerator for 15 min;
  5. Pour methanol (organic solvents), into a pot and discard properly.
    1. ** For fibers, remove one by one and position in the respective location of the well of the lid. Pour the plate and return them;

      (Work in the chapel)

      1. Take the amount in a beaker (100ml);
      2. Discard in an ice cream jar;
      3. In the end, discard everything, including the unused, in the discard pot.
    2. Squeeze the plate against a paper and let it face up;
    3. Add 2ml violet crystal and leave for 20 min;
    4. Pour the violet crystal and gently wash the plate with milli-Q water;
      1. ** For fibers, remove one by one and position in the respective location of the well on the lid. Wash the plate and then the fiber one by one, returning them to their respective wells.
    5. Remove excess water with absorbent paper;
    6. Add 2ml of 30% acetic acid (total 32ml / plate), and homogenize each well;
    7. Transfer 125ul from each well to a flat plate with 96 bottom wells, for the Abs reading at 590nm.

Coconut fiber production

NOTE: We bought the green coconut in the market.

Materials:

  • Green coconut
  • Knife
  • Beakers
  • Blender
  • Sieve
  • Distilled water
  • Use the knife to open a window in the coconut.
  • Remove the coconut water.
  • Cut the coconut into some slices using the knife.
  • Remove epicarp, endocarp and edible part.
  • Insert pieces of the mesocarp of about 5g in the blender and add distilled water until it covers them.
  • Beat in blender for about 2 min.
  • Remove from the blender and, using a sieve, wash the fiber with distilled water.
  • Place the washed fiber in beakers and allow it to dry in a humid incubator at 37ºC, for about 8h.

Biofilm quantification using crystal violet 1.0

Note: 3 days protocol

Materials:

  • Acid acetic 30%
  • Medium LB
  • Medium LB + glicose
  • Violet Crystal 0,1 % in milli-Q water
  • 96-well plates

Day one: Preparing solutions and pre-inoculum.

  1. Prepare all medium and solutions to prepare pre-inoculum;
    1. Acid acetic solution 30%;
    2. Medium LB (pre-inoculum);
    3. Medium LB and LB + glicose for biofilm growth.
  2. Autoclave all mediums;
  3. Prepare 5 mL pre-inoculum in medium LB and incubate overnight (12h - 18h) in 37 ºC;

Day two: Biofilm quantification using violet crystal.

  1. Dilute overnight culture in 5:100 (medium LB);
  2. Add 100uL in each well of the plate;
  3. Cultivate static for at least 24h in 37 ºC.

Day Three: Measurements.

  1. Discard all cells turning the 96-well plate in a bowl with sodium hypochlorite using water to clean the wells;
  2. Add 125uL of violet crystal 0,1% in each well;
  3. Incubate for 10-15 minutes in ambient temperature
  4. Wash 3 or 4 times with water to remove the excess
  5. Let dry for a few hours or overnight

    Quantification:

    1. Add 125uL of acid acetic 30% in each well
    2. Incubate 10-15 minutes in ambient temperature
    3. Transfer 125uL for a new 96-well plate with flat bottom
    4. Measure absorbance in 550nm in the spectrophotometer using acid acetic as blank

PCR - Phusion High-Fidelity DNA Polymerase

The protocol used for this PCR can be found in the official NEB website and followed according to manufacturer's instructions.

This part of our project revolves around two different circuits: the first one containing the Secretion System Type I, and the second containing our new part, MermAID. In order to assemble the synthetic DNA we ordered from TwistBioscience, we had to perform a PRC for each fragment, removing the adaptors from the edges. We used Q5 Polymerase enzyme. The following protocol was used: PCR - Q5 High-Fidelity 2X Master Mix (NEB).

Also, since our primers had different annealing temperatures, we used a thermocycler with a temperature gradient shown in the images below:

Bio-Rad T100 Thermocycler
Bio-Rad T100 Thermocycler
Bio-Rad Horizontal Electrophoresis Systems
Bio-Rad Horizontal Electrophoresis Systems

We then ran an agarose gel and purified the product. We measured the final concentration using NanoDrop. The following protocols were used: Preparation 0.8% agarose gel, Purification of PCR product from agarose gel and Concentration measurement - Nanodrop.

In order, A1, A2, A3, A4 represent the fragments from MermAID and M1, M2, M3, M4, M5, M6 represent each fragment from the secretion system. A2 and A3 were expected to have around 0.3 kb, and the others around 1.0 kb:

Agarose gel - PCR product, amplification of our synthetic DNA fragments
Agarose gel - PCR product, amplification of A1, A2 and A3 respectively

From the picture next to, we can see that most fragments were correctly amplified, with the exception of the first three, that did not show clear bands. This could have happened as a result of the complexity of these DNA fragments, since they have high CG content and could possibly be forming secondary structures. To eliminate the possibility that it was due to human error, we repeated the PCR of these three fragments with the exact conditions previously used.

As seen, they continued to present a different behaviour from what was expected. Even though we can see a define line where our fragments were supposed to be, it still differs from the other ones, that were in much higher quantity and did not have the visible drag present in these ones. The next step would be to assemble all fragments using Gibson Assembly, but in the light of the result, we decided to perform another PCR, to try and fix these three fragments. In this new attempt, we added 3% DMSO.

Agarose gel - PCR product, amplification of A1, A2 and A3, with 3% DMSO

From the picture above we can see that DMSO wasn’t effective as we hoped. We then proceeded to test PCR in different temperatures around primers Tm for each of these three fragments, without DMSO.

Agarose gel - PCR product, amplification of A1, A2 and A3, with different annealing temperatures
Agarose gel - PCR product, amplification of A1, A2 and A3, with different annealing temperatures

After trying everything we could think of and failing to obtain the desired result, we decided to give one more try and change the enzyme used in PCR, from Q5 to Phusion, with and without DMSO 3%:

This was the best PCR product we had so far, so we decided to proceed with the experiment and ran another gel to purify the product:

Agarose gel - PCR product, amplification of A1, A2 and A3, with and without DMSO 3%, respectively
Agarose gel - PCR product, amplification of A1, A2 and A3, with and without DMSO 3%, respectively
Team member cutting agarose gel to purify PCR product

We measured all fragments concentrations using NanoDrop:

SAMPLE

CONCENTRATION (ng/µL)
A1 no DMSO 20.65
A2 with DMSO 9.7
A3 with DMSO 11.7
A4 37.8
M1 11.15
M2 19.8
M3 21.7
M4 25.2
M5 29.8
M6 23.37

In the meantime, we designed and ordered primers that would allow us to amplify the vectors in which our circuits would be inserted. Unfortunately, our order got delayed by two weeks, preventing us to advance with the assembly for a couple of weeks.

When the primers finally arrived, we amplified the vectors pSB1K3, and pACYCDuet. At this point, we are interested in the pSB1K3 vector, which will be used in both assemblies, MermAID and Secretion System. It will be crucial to remember where we got this pSB1K3 vector. It was amplified from the iGEM DNA Kit (BBa_J04450), and it consists of a part containing pSB1K3 + RFP. We designed the primers in a way to open the vector and remove the red fluorescent protein. The expected band was around 2.0 kb:

Agarose gel - PCR product, amplification of pACYCDuet for M1, pSB1K3 and pACYCDuet for MerChina with and without DMSO, respectively

We finally have a result we can work with! With this last agarose gel, we had what was necessary to assemble all fragments. So we purified them, and measured their concentration using NanoDrop. The next step was to unite them using Gibson Method, and transform in DH5-Alpha according to the protocols NEBuilder HiFi DNA Assembly Reaction Protocol and Bacterial transformation 1.

In order to do so, we had to do some math. The protocol for the Assembly MasterMix we used required specifics amounts of DNA according to the number of fragments assembled. Let’s spare you from all the numbers and skip to the part we finally had both circuits assembled into the vectors. We transformed them into DH5-Alpha.

From left to right: plate with MermAID + pSB1K3 transformed in DH5alpha and Secretion System + pSB1K3 transformed into DH5alpha

If we zoom in, we can clearly see that there are two different types of colony in both plates, white and pink:

But wait a minute… how can it be? After some deliberation, we came up with a possible explanation. Remember a few paragraphs above, where we mentioned it would be crucial to know the origin of the vector? Well, our theory was that even after we amplified and purified our DNA, some vectors didn’t receive our insert, possibly due to presence of prefix and suffix on the edges of vector backbone, that are the same of RFP, so it could end up closing again with RFP protein. Therefore, the pink colonies would be the negative control, and the white ones would be the ones we are interested in. In order to confirm this theory, we performed a Colony PCR for each plate, one for the MermAID and other for Secretion System. We followed the protocol PCR - Phusion High-Fidelity DNA Polymerase.

To confirm that the cells have our insert, we selected six white colonies from each plate (from 2 to 7) and one pink (8):

Legenda
Legenda

We expected to see M samples from 2 to 7 with a band around 5.6 kb, and A samples from 2 to 7 with a band around 2.2 kb. M8 and A8 were supposed to have a band with a size corresponding to RFP protein, around 1.0 kb:

With this picture we can see a band corresponding 1.0 kb in A8 and M8 samples, as was supposed to, according to the previous analyses. With regard to the other M samples, it’s clear that the 5.6 kb band was not present in any of them. In what concerns A samples, we see a lot of bands with differents sizes from the expected. Two intense bands drew our attention, both at same height on the gel, between 2.0 kb and 3.0 kb, relative to sample A3 and A6. Even though these bands sizes are as expected, the samples also have other defined bands at different sizes.

Despite the fact that A3 and A6 were not 100% pure in respect to our insert band, they confirmed its presence. But it’s also interesting to note that not all colonies present in the plates were transformed with a vector containing our inserts.

This was a milestone in our experiments, because with this gel we confirmed that our Gibson Assembly reaction worked, and that our insert was indeed inside the vector. Having this confirmation enabled us to proceed with the planned experiments, using colonies A3 and A6.

After accomplishing the constructions MermAID and Secretion System, the next step consists in creating a biological circuit with our chimeric protein Iara-alpha under the regulation of a easily induced promoter.

Instead of using the original idea of Mer promoter, induced by the presence of Hg inside the cell, we thought of the importance of having an induced promoter, which allows us to yield the protein expression at any time, not only under a mercury stress condition. Using the Mer promoter, the functionality relies on the bacteria having a proper transport system to bring Hg inside the cell.

Therefore, the Mer promoter should be switched by the lac promoter, easily induced by adding IPTG. This line of thinking also goes hand in hand with our application proposal of a functional biofilter.

In order to do it, besides a promoter alteration, the initial idea of using an iGEM vector to insert the chimera Iara-alpha, had to change. Due to the shared ColE1 family replication origin between then, we opted for the construction Iara-alpha + pACYCDuet, a plasmid compatible with iGEM vectors for a future co-transformation with the Secretion System circuit. The pACYCDuet works under the lac promoter and has resistance for chloramphenicol. Another advantage is the possibility of adding a His-Tag in one of the Iara-alpha extremities, allowing it’s purification by affinity chromatography.

In the wet lab, after obtaining the complete assembly of MermAID in pSB1K3, we proceeded with PCR to amplify the region corresponding to our protein Iara-alpha.

After executing Gibson Assembly, transformation of the construction Iara-alpha + pACYCDuet and the attempt to confirm it’s product by performing a colony PCR, we chose two samples - A3 and A6 - to prepare pre inoculum for future Iara-alpha amplification. We also made a plate with one colony of each sample, to work always with the same cells.

Unfortunately, the A3 pre inoculum didn’t grow, so we went forward only with A6, concentration of 114.13 ng/uL, performing PCR with Q5 Polymerase enzyme. In the verification agarose gel we weren’t able to see any band, so we came to the conclusion that our chimera Iara-alpha , approximately 1.0 kb, had not been amplified.

Agarose gel - The three strongest bands of molecular marker corresponds to 3.0 kb, 1.0 kb and 0.5 kb respectively.

To get our chimera, we tried another round of PCR, with a few modifications in order to improve the chances of amplifying our sequence. PCR reactions were completed with the addition of DMSO, BSA and glicerol, in different combinations, using Phusion Polymerase. Total reaction volume: 50 uL, water volume added to fulfill the reaction total volume.

SAMPLE

DMSO (%)

BSA (µg)

GLICEROL %

DNA TEMPLATE (ng)
1A 0 - - 100
1B 3 - - 100
1C 5 - - 100
2 3 50 - 200
3 3 50 3.125 200
Agarose gel - second round PCR: wells corresponding to 1A, 1B, 1C, 2 and 3. The three strongest bands of molecular marker corresponds to 3.0 kb, 1.0 kb and 0.5 kb respectively.

We see a weak band at around 1.0 kb, corresponding to what was expected of our protein Iara-alpha. Unfortunately, after miniprep purification the final concentrations were very low:

SAMPLE

CONCENTRATION (ng/µL)
1A 3.0
1B 3.2
1C 3.2
2 3.3
3 3.6

Despite the low concentrations obtained, we chose the highest, sample from colony A3. To unite the chimera and it’s backbone, pACYCDuet was linearized and properly amplified to contain overlaps on it’s extremities to fit the Iara-alpha:

Agarose gel - PCR product, amplification of pACYCDuet for M1, pSB1K3 and pACYCDuet for MerChina with and without DMSO, respectively.

We were now ready to proceed with the Gibson Assembly method. The reaction product was transformed into E. coli DH5alpha, but after 24 hours, the plate didn’t show any colonies. Consequently, the reaction wasn’t successful and we didn’t get our desired construction.

Plate of transformed E. coli DH5alpha with Gibson Assembly product.

In front of the result and reviewing the calculations for the assembly reaction, we decided to perform the same reaction again, altering parameters. The total volume was changed from 20 uL up to 60 uL, so that we could have more of our interest DNA in order to form the wanted construction. Unluckily, the plate was empty, without transformed cells.

Plate of transformed E. coli DH5alpha with Gibson assembly product, second reaction with total volume of 60 uL.

In order to advance, we decide to take a step back and prepare another pre inoculum from the colony A3. The plasmid was extracted through miniprep, with concentration of 187.3 ng/uL. Another PCR was performed using Phusion Polymerase, altering reaction parameters:

SAMPLE

DMSO (%)

BSA (µg)

GLICEROL %

DNA TEMPLATE (ng)
α 5 - - 100
β 3 50 - 100
Υ 3 50 5 200
Agarose gel PCR of chimera with Phusion Pol: wells corresponding to &alpha, &beta, and &Upsilon. The three strongest bands of molecular marker corresponds to 3 kb, 1 kb and 0.5 kb respectively.

The expected bands were supposed to be around 1.0 kb, but we can clearly see from the picture above that something along the way went wrong, and we ended up having three crear bands around 3.0 kb.

At this point of the project, considering the amount of time left, we decided to give Iara-alpha a last chance of being amplified. We agreed in go backwards a few steps and start pre inoculum with 5 different colonies of the white-pink plate: A10, A11, A12, A13 and A14 besides the previous A3 and A6.

White-pink plate: MermAID + pSB1K3 transformed into DH5alpha.

After plasmidial extraction and purification we obtained the following concentrations:

SAMPLE

CONCENTRATION (ng/µL)
A10 25.0
A11 28.11
A14 31.8
A3 35.0
A6 22.3

The last PCR was performed with the optimized conditions so far: working with Phusion Polymerase, DMSO 3%, BSA and glicerol.

Agarose gel - The three strongest bands of molecular marker corresponds to 3.0 kb, 1.0 kb and 0.5 kb respectively.

This last agarose gel showed that even after trying many different approaches, we couldn’t amplify our protein Iara-alpha, and therefore we were forced to drop this line of experiments. We decided to readapt our plan, and go back to working with MermAID, even if the planned experiments and tests weren’t ideal for this specific synthetic system.

Hg Disc Diffusion Test

This step of the experiments intends to assess the survival capacity of our transformed bacteria containing the insert MermAID. We expect that due to it’s Iara-alpha expression and consequently binding to Hg, the bacteria would be able to survive more in a mercury stressful environment than a non-transformed one. With this data, our bacterial growth shows up higher than a wild type E. coli DH5alpha, a consistent explanation would rely in the functionality of our biosynthetic circuit, allowing inference about its effectiveness.

The growth is going to be compared through the bacteria presence or not, around a diffusion disc with mercury solution of the salt HgCl2. The halo size and the proximity to the disc indicates how resistant the E. coli DH5alpha culture was, or, in a indirect way, if the MermAID works or not.

We designed the experiment performing it using solid LB agar plates, each one containing duplicate of a paper filter disc wetted with mercury solution, following the protocol “Hg halo disc diffusion test”. The tested working concentrations of HgCl2 were 0 ug/mL, 20 ug/mL, 200 ug/mL and 20 mg/mL, a concentration per plate. On the disc diffusion control, with zero Hg, water was applied instead of mercury solution.

Hg CONCENTRATION (µg/mL)

Hg MASS (µg)
20 0.2
200 2
20000 200
Quantities of Hg in each concentration for the respective filter paper circle ( Diameter: 12 mm)

In order to do it, we used 5 selected colonies of the transformation white-pink MermAID plate: A10, A11, A12, A13 and A14. Besides that, we also did plates containing the non transformed E. coli DH5alpha and a contamination control plates without cells.

In summary, the performed experiment involved 28 plates, a set of three Hg concentrations plus water control for each selected colony. The solid cultures were incubated for 24 hours, at 37ºC.

Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.
Hg disc diffusion test plates, each column corresponds to a colony, with the concentrations range from 20 mg/uL away down to zero.

Look to the plate results, was observed that non transformed E. coli DH5alpha, our biosynthetic circuit functionality control, grew in all 3 Hg concentrations, which wasn’t expected, since they do not have the insert that would confer them resistance to Hg, which is toxic and theoretically causes its death. Another aspect observed was that every plate with 20 mg/mL Hg concentration presented a halo zone of the same size, while in the others lower concentrations the bacterial culture spread through all plate. It means that for some reason our bacteria performed just like a non transformed one, having the same Hg radius of inhibition.

The plates analysis lead us to the formulation of 3 hypothesis to explain what happened:

  1. The Hg quantity applied in each filter paper circle, 10 uL, wasn’t enough to cover the hole plate;
  2. The 200 ug/mL and 20 ug/mL Hg concentrations are small and allow non transformed bacterias to survive;
  3. The bacteria does not contain our insert.

    In front of this results and hypothesis, we decided to repeat the experiment, changing some parameters. We tested the size of the filter paper circle, to guarantee that the volume used was enough to wet, but not to soak it. With this, we decided that the initial size was perfect. Also applied Hg solution in a more systematic way to guarantee the same waiting time and to establish a more solid replicable method.

    In the following attempt, to ensure formation of different halo zones sizes, we added three different Hg concentrations to the previous ones: 2000 ug/mL, 10000 ug/mL, 40000 ug/mL and excluded the lower one of 20 ug/mL. This way, we could analyse if there’s a minimum Hg concentration that belows it the natural cells can survive without trouble and beyond our circuit is necessary for bacterial survival. The last variable we changed was the colony used, the A3. More details of why can be found in Lab Notes, date 10.19.2019.

    The plates were made in triplicate and each large plate was divided in six, so that every plates had all Hg concentrations. The indicated control is the non transformed E. coli DH5alpha:

    Hg disc diffusion test plates, with concentrations ranging from zero to 40 mg/uL.

    In order to properly analyse the relation between Hg presence and cell growth the halos were measured using ImageJ, to maximize measurements precision the area of each halo was characterized 5 times, we then took its average value and compared obtained values in each replicate. The next step is to plot histograms with the average values of our experiment replicates vs. the control culture, comparing cell behaviour around the Hg contaminated zones. We expect to find smaller halos, i.e. smaller radii of death zones, in transformed cells due to their capacity to capture Hg which is believed to enhance cells chance to survive in the hostile environment. The table below shows the measured radii for each plate and then the average values for each concentration.

    The graph displayed exhibits the constructed histogram of average radii vs. control culture. In this case, the culture was incubated for 12h and the results indicate that transformed cells were able to survive in Hg contaminated zones better than non transformed cells, in particular for higher Hg concentrations which cells were not expected to survive at all.

    When comparing replicates behaviour, we are able to identify that transformed cells might have an optimal Hg concentration to outgrow non transformed cells around 1000 ug/mL. The results shown above indicate that the are cells are suitable for our projects interest due to the results obtained for concentrations above 20 000 ug/mL.

    Graphic: Measurements of halos from mercury stained colonies after 24h.

    We measured the radius again after 24h. We can note by the graphic that the radius of transformed cells increased a little if compared to 12h, and the non-transformed cells didn’t show any difference between the previous measurement. It can indicate that our bacteria is more resistant in different concentrations of Hg, but they eventually reach a threshold point, and also start to die which, increases the radius. Although the radius had increased in 24h for transformed cells, it is still smaller than non-transformed cells, which can support our hypothesis that our circuit helped to improve mercury resistance.

    Hg growth curve

    This experiment consists in mapping the bacterial growth of E. coli DH5alpha containing our biosynthetic circuit versus non transformed ones, in medium with different mercury concentrations. This way, we would be able to compare the behaviour of our transformed cells in the presence of Hg, with non transformed ones. With this we hoped to demonstrate that cells containing our biological circuit were more fit to survive in and medium containing Hg, and therefore prove that not only our protein was being expressed, but also that it was properly exerting its function to capture mercury.

    We worked with some of the previous Hg concentrations used in “” HALO DEATH and another ones between (0-200) ug/mL: 0 ug/mL, 7.5 ug/mL (same concentration of team UFAM 2016 used), 20 ug/mL, 200 ug/mL, 2000 ug/mL. Cells were incubated at 37°C under agitation. The measurements were made every 30 minutes starting with OD600 = 0,05.

    For the experiment, we transformed into E. Coli DH5- the metal binding chimera (Iara-) and the GGDEF domain-containing protein (Q9X2A8), both regulated by same Mer promoter. The insert was into the pBS1K3 vector.

    We started the test with a low optical density and keep measuring it every 30 minutes within 8 hours (for the transformed bacteria) and within 6 hours (for the unmodified strain). The bacteria was growth in culture medium Luria-Bertani (LB) containing different concentrations of mercury such as 0, 7,5, and 20 µg/ml. For the transformated strain we also made the experiment with high mercury concentrations such as 200 and 2000 µg/ml. The results are shown in the images below.

    As can be seen on the graphs above, in the experiment performed with 0 µg/ml the insert-containing bacteria had a slower growth compared to the unmodified strain. This behavior may be due to increased metabolic expenses of transformed bacteria to express the synthetic proteins. Moreover, the transformed bactéria was able to grow in culture medium containing 7,5 µg/ml while the unmodified strain failed to grow in this concentration. Thus we can infer that our metal pickup chimera, Iara-α, gave to the bacteria a greater resistance to mercury, making it able to survive in environments with the concentration of Hg tested. Also, this is an evidence that Iara-α is been expressed and it is working as desired. In higher concentrations neither the engineered bacteria nor the unmodified one survived, since mercury ions are highly toxic this result was expected.

    These experiments also showed that our MermAID works even without the Secretion System, which would be responsible to lead our protein out of the cell. Instead of binding to our protein outside the cell, the mercury penetrates the cell, where our protein binds to it. The downside is that bioaccumulation leads to cell death for high mercury concentrations.

    This behaviour is evidenced in the growth curves, because even though transformed cells survived longer for low concentrations, when compared to the non transformed ones, as we increased the Hg concentration, they could have reached a threshold, saturating the amount of protein available to capture the mercury, and consequently causing cell death.

One part of our project consists in the expression of a di-guanylate cyclase (DGC), a gene that contain a GGDEF domain responsible for induction of biofilm formation. In order to find the best combination of elements to build a functional and efficient circuit, that is, a circuit that is able to produce biofilm, we performed a range of tests.

Testing the best chassi for biofilm formation

To begin, we intended to test the best chassi for our DGCs, in order to verify if one of the available bacterial strain in our laboratory naturally produces biofilm. The studied parameters were: time and culture medium. With regard to time, we assessed if 24 hours incubation time was enough for natural biofilm formation. And for the medium, we decided to investigate the influence of glucose presence, testing two types: LB with and without it.

96-well plate set up

With this test, we hoped to see a different behavior in columns 3 and 6, since they were the ones containing bacterias. This difference is shown through distinct stain levels when we mark the biofilm with violet crystal. This was made according to Biofilm quantification using crystal violet 1.0 protocol.

Well plate 24 hours.

As expected, the wells of columns 3 and 6 appeared to be more stained than the controls, indicating the presence of biofilm. However, some of the controls with glucose were stained (4B, 4D, 4E, 5D, 5E, 5F and 5G) - showing a possible contamination. Absorbance was measured in a spectrophotometer and the data treated with software SpectraMax 6.4:

Results indicate a possible contamination of the plate, due to absorbance levels measured especially in the controls 4 and 5. Apart from that, the columns 3 and 6 showed biofilm production, and between the two mediums, LB+glucose appeared to be the better one. This test evidenced that we had to make some changes in our initial protocol: we added a step to normalize optical density of the cultures (DO), and had find a more efficient way to clean the plate after staining it with violet crystal, since this could mislead us to false conclusions about the biofilm formation.

Testing production of biofilm for 24 and 48 hours with static and agitation bacterial growth

The next step in the experiment is to study and characterize our DGCs. In order to do so, we repeated the violet crystal staining test on a 96 well-plate, but this time we used pETSUMO vectors with our respective DGCs as inserts. It’s relevant to note that pETSUMO vector is induced by IPTG and has kanamycin resistance.

We obtained all miniprep vectors by transforming them in DH5alpha for cloning, and then in E. coli BL21 for expression, both transformed with the following backbones:

  • pETSUMO::YdeH (BBa_K1019004)
  • pETSUMO:: wspr (BBa_K3280001)
  • pETSUMO:: yddv (BBa_K748003)
  • pETSUMO::empty
DH5alpha transformed with pETSUMO::DGC’s - pETSUMO::wspR
DH5alpha transformed with pETSUMO::DGC’s - pETSUMO::ydeH

Once we had all plates, we were ready to proceed with the biofilm formation test.

The 96-well plate map used can be seen below:

<

IPTG Concentration

1

2

3

4

5

6

7

8

9

10

11

12
A Without IPTG Control YdeHwspr yddv
B 0.01 mM
C 0.1 mM
D 0.5 mM
E 1 mM
F LB+kanamycin
G
H

All the plates had DO of cultures normalized, so they all start approximately with the same number of cells. we tested static growth for 24 hours and two replicates.

We expected to see M samples from 2 to 7 with a band around 5.6 kb, and A samples from 2 to 7 with a band around 2.2 kb. M8 and A8 were supposed to have a band with a size corresponding to RFP protein, around 1.0 kb:

First 96-well plate 24 hours: where the cultures grow. Second plate 96-well plate 24 hours: where the measures are done.

Quantification of biofilm with violet crystal, according to the literature, are supposed to show purple stains visible to the naked eye. Looking at the plates, we didn’t see any stained wells and no difference between them, which diverged from literature data.

Graphics below are from two tests made in different plates and neither indicated biofilm production:

With the absorbance values we noticed that controls exhibited similar numbers to the plates with transformed cells, which wasn’t expected. In the light of these results, we decided we had to readapt our protocol once more. So on the next round of tests, we tried the following protocol: Biofilm quantification using violet crystal.

Before continuing with testing, we decided to review all the past steps and we found problems in the begging, with the plating of BL21 in a wrong concentration of antibiotic. This could possible generate growth of unwanted bacterial colonies, explaining the negative results. So we transformed all DGC’s again and restarted the experiments.

BL21 transformed and marked colonies to do another plate.

After transformation we selected one colony of each one, pre-inoculated and made another plate that has bacterias of the same origin. Before this point we started an adapted protocol with the news transformed bacterias.

We changed some important points of the previous protocol:

  • 24-well plates for increase volume of medium to guarantee the growth of bacterias;
  • After pre-inoculum bacterias were transferred to SOC Medium with initial DO=0,2 and let them grow for 4 hours before being added in plate;
  • Wash of the 24-well plate with PBS;
  • Addition of a step with methanol for biofilm fixation;
  • Growth with agitation.

DO were measured at 600 nm and diluted twice in SOC Medium:

SAMPLE

DO DILUTED 2x

REAL DO
yddv 0.65 1.3
wspr 0.5 1.0
YdeH 0.4 0.8
Control 0.4 0.8

This measures enable DO normalization in 24-well plates YdeH to all the bacterias start at the same point. Also this result presented that YdeH has lowest growth between others DGCs and yddv the higher.

We decided to run a pilot-test, in order to see if our protocol changes would work. This is our first construction for tests and IPTG concentration was the same for all the wells (1 mM):

1

2

3

4

5

6
A Blank YdeH Blank
B yddv
C Control
D wspr
24 hours 24-well plate after acetic acid addition.

With this plate we can visually infer that wspr showed the lowest biofilm growth, and YdeH the highest. To confirm this, we measured the absorbance at 590nm for each well:

DGCs.

This graphic comproves our visual analyses, showing that YdeH DGC bar is higher than the others, and wspr lower. 24-well plate compared to 96-well plate showed more conclusive and consistent results and no contamination, since coloration of each line is more homogeneous.

We then ran our official tests using the same protocol used in the pilot-test. To decrease bar error, we added one more replicate on the plate and different incubation times. We also added a plate to grow static, for future comparison between methods. We maintained 1 mM IPTG concentration for all wells.

1

2

3

4

5

6
A Blank Controle
B wspr
C YdeH
D yddv

Our first try went wrong, the plate didn’t show any signs of biofilm formation.

24-well plate after 24 hours in agitation, showing a transparent medium with sedimented bacterias.

We later realised our mistake in preparing SOC medium, we used the wrong ratio of reagents. We discarded the plates because after 24 hours the cell were dead, probably due to poor nutrients supply. So we repeated all the steps with the correct medium and obtained the following results:

24-well plate after 24 hours in agitation, showing a transparent medium with sedimented bacterias.

Plates showed a clear difference to the naked eye, as the intense stained on the last line of the right plate on the image. Experiment with agitation has contamination on the blank, possibly due to movimentation of the culture medium inside incubator that could overflow from the well. Less homogeneity is also observed in the second plate, showing that experiment with agitation could provide more unstable conditions.

Each column is the mean value of a sample with 5 identical experiments made in the same 24-well plate, with the exception of the static SOC medium which had 1 of its results masked due to their dissonance from the rest, implying external contamination.

As it can be seen from graphic, all DGCs showed a higher biofilm production in a non-static condition, with the exception of the bacteria transformed with WspR, which is coherent with other biofilm growth experiments made with E. coli. The DGC that produced higher amounts of biofilm under agitation was clearly YddV, with an absorption rate more than 6 times higher than the second biggest producer, agitated YdeH.

It’s interesting to note that a certain quantity of biofilm was expected for the control, pETSUMO without DGC insert, since our bacteria naturally produces low rates of biofilm.

24-well plate after 48 hours in agitation.
Once again, 5 replicates were made, with dissonant results masked

The 48h static plate showed lower biofilm formation if compared to the 24h static plate. This could indicate a natural tendency for the E. coli to not maintain the small amount of biofilm that they produced under static conditions, since forming this matrix it’s a costly metabolic function.

As for the 48h agitated plate, the absorption rate didn’t change significantly for the bacteria transformed with YddV and WspR, indicating a possible saturation in the amount of biofilm maintained by those cells at approximately 3 Abs. The cells transformed with YdeH showed the biggest change in the absorption rate from 24 hours to 48 hours, indicating a preference for a longer period of incubation under agitated conditions.

The agitated WspR showed a very heterogeneous growth similar to the agitated YdeH in the 24 hour plate. We can infer that this is a form of response from this DGC and that these bacterias would reach a higher level of absorbance if a longer incubation time was tested.

The same results can be presented in a different manner to evaluate the changes happening from 24 hours to 48, by plotting the change in absorption against time:

Once again, 5 replicates were made, with dissonant results masked
Once again, 5 replicates were made, with dissonant results masked

To conclude, we observed a substantial increase in biofilm formation for plates incubated under agitation, as well as for longer periods of time. We also noted that under static conditions, cells tend to considerably decrease biofilm production. Moreover, the DGC that showed the fastest response in the matter of biofilm production was YddV, which managed to saturate the acetic acid solution with biofilm in at least 24 hours. The second fastest response was from the DGC YdeH, reaching the maximum absorbance level in 48 hours. The agitated wells containing wspR manage to increase its biofilm production in the span of 24 hours, but did not reach the same high levels as YdeH and YddV, a longer timespan is necessary to evaluate wspr biofilm production.

Verification of biofilm adherence in coconut fiber

Our final system should include the presence of coconut fiber in the biofilter that works as substrate to adhesion of biofilm. This experiment has the idea to simulate the real situation and verify the percentage of biofilm on the fiber that was immersed in a culture of transformed bacterias.

Previously in our saga, we obtained that growth with agitation showed the best results and increase of time, also exhibited increase of biofilm production. Therefore, we planned 24 well plates for 48 and 72 hours with agitation following protocol 21 including an extra part for the coconut fiber.

We used a coconut fiber that was characterized by the protocolo 23. So to put in the 24 well plate we did little balls of them with weight 60 mg.

Preparation of the plates with coconut fiber.
Preparation of the plates with coconut fiber.

We used the same plates of Dh5alpha transformed with pETSUMO ::: DGCs and SOC medium to do this experiment. We did 4 blanks, and 3 replicates of each sample:

  • pETSUMO::YdeH (BBa_K1019004) - YdeH
  • pETSUMO:: wspr (BBa_K3280001) - wspr
  • pETSUMO:: yddv (BBa_K748003) - YddV
  • pETSUMO::empty - Control
  • Blank - SOC medium without bacteria

1

2

3

4
A Blank Controle
B wspr
C YdeH
D yddv

After 48h and 72h we separated the fiber and put in another plate (let’s call plate F). We followed the procediments from the protocol: NUMBER 21.

After the part using violet crystal in our plates we could see qualitatively some differences between the amount of biofilm in the plate and in the fiber.

On the image below we can notice an intense color of crystal violet on the wells by naked eyes, probably due the high absorption of the fiber:

Plate after 48h - plate on the left is the fiber one and plate on the right is the plate where the fiber was before fixation with methanol.
Fiber after violet crystal protocol (72h)/ Plate where the fiber was after violet crystal protocol (72h)

Analysing after addiction acid acetic to our plate we noticed that some samples of control and blanks were as violet as the DGCs, it probably is a sign for contamination between the wells.

Once we finished preparing the plates we measured them by absorbance with 590nm.

Experiment of biofilm formation with fiber in 48h.

Comparing with previous experiments, there is a possibly of biofilm adhesion on coconut fiber given it’s decreased stained in the plate reading. We measured absorbance of crystal violet in both components: fiber and plate, to confirmate the distribution of biofilm growth. Our plan was use LB broth as a normalization of the experiment but after measures we realized that could possibly occurred a saturation on absorbance value that didn’t surpassed 3,3.

Absorbance of experiment biofilm formation with fibers in 72h.

We obtained a similar result for 72 hours well plate for the fibers and an increase of growth in the plate compared with 48 hours. Limit of absorbance remained the same, reinforcing the idea of a saturated solution of crystal violet, preventing our initial comparison plan.

So with this experiment we tried to measure the concentration of biofilm with coconut fiber. But we noticed that our results had a methodological problem because although we realized that there was a biofilm, with 48h and 72h of growth we reached the acid acetic and violet crystal saturation and we couldn’t conclude how much biofilm grown up adhered.

FABRICATION

To reduce filter production costs, we decided to produce coconut fiber through a green coconut.

Coconut fiber production protocol

We drilled a hole in the green coconut and removed the water. After the coconut is empty, we break the coconut, we remove the epicarp (smooth epidermis/coconut shell), the endocarp (a hard layer that surrounds the edible part) and the edible part of the fruit.

That left only the mesocarp (fiber bundle), we separated it into some "handfuls" of fiber and put it together with distilled water in the blender. After beating for a few minutes, we remove the fiber and wash it with flowing water.

The fiber, already beaten and washed, was placed in beakers and it dried in a humid incubator at 37ºC, for about 8h.

CHARACTERIZATION

Absorption of water protocol

The coconut fiber water absorption test was performed so that we could better plan the volume of fiber and the water flow we should be using in the filter.

We weighed 1g of coconut fiber, which was placed in 50mL beakers and we added different volumes of distilled water until complete material humidification (overnight), which was determined by visual observation if there is visible water in aqueous phase.

The table below shows the volumes used and whether or not it has been absorbed.

Volume of water (mL)

Absorbed?
2.5 yes
3.5 no
5.0 no
10.0 no
20.0 no

As can be seen from the table, of all tested volumes only 2.5mL was fully absorbed by the fiber after the analyzed period. Therefore, we know that the max absorption of our fiber is 2.5mL/g.

Fourier-transform infrared spectroscopy - FTIR

The FTIR equipment was used as part of the characterization of our fiber since it was possible to determine the presence of cellulose in our coconut fiber. This is important because as our mercury-collecting chimera has a cellulose-binding tag (inserted by us into the genetic circuit), the protein can bind to both the bacterial biofilm matrix and coconut fiber (if it had a significant percentage of cellulose).

We used the FTIR equipment from the laboratory of the Nanotechnology and Nanotoxicology Group (GNano) of our Institute. With the help of postdoctoral student Bianca Martins Estevão, we made KBr pellets with the green coconut fiber powder, and with these pellets, we were able to collect the FTIR spectrum of our fiber.

Comparative analysis of the spectrum with the literature [9,10,11] shows that there are bands attributed to hydroxyl groups (O-H cellulosic stretch) at 3430 cm-1, axial deformation of C-H groups at 2910 cm-1, angular deformation of C-H groups 1375 cm-1, angular deformation of primary alcohol C-O bonds at 1167 cm-1, C-O-C bond absorption band, representing pyranose ring vibration at 1050 cm-1 and β-glycosidic bonds between glycan units in 901 cm-1, characteristics of cellulose. The band at 1745 cm-1 refers to acetate group residues from hemicellulose.

With this result, we could conclude that coconut fiber could possibly serve as a good substrate for the cellulose binding anchor of our metal-collecting chimera.

Coconut fiber cellulose FTIR spectrum after being ball milled.

Due to the high cost of mercury concentration measurements and the need of relatively high volumes of mercury solutions that would be required for laboratories to provide reasonable results using traditional methods of analysis, as ICP-OES [4], our team had to work on alternative methods of metal quantification or even semi-quantitative analysis procedures.

For this, we try a semi-quantitative approach using Laser Induced-Breakdown Spectroscopy (LIBS). The available equipment to carry on this analysis belongs to Empresa Brasileira de Pesquisa Agropecuária (Embrapa), São Carlos unit [5].

Using several mercury concentrations in solutions we performed the calibration. We then dripped 10 uL of our solution on different KBr pads and left it drying for a few minutes before putting it to the sample holder.

For every laser hit, a spectral emission curve was generated, and after 100 laser hits we were able to generate an average emission spectrum of samples.

We have previously searched for Hg peak emission wavelength, found it at 194.15 nm and 253.65 nm, thus when treating the collected data we could analyse with higher precision the regions around these peaks, proceeding to data normalization right after.

Finally, we plotted the graphs shown below, they show the emission spectrum and a bar graph of normalized area of each peak emission of both of the Hg emission peaks.

The KBr powder to make the KBr pellet was 98% pure and there was a low concentration of Fe II. Fe II has a characteristic emission peak that is very close to one of the mercury emission peaks at 253.65nm.

When analyzing the emission spectrum of KBr pellets with and without mercury solution, we found no significant difference in the peak relative to the emission of 253.65nm, probably due to Fe II emission present in the KBr pellet composition. Thus, we chose to take into account only the part of the spectrum referring to the 194.15nm peak.

Graph 1 shows the average emission peak of 194.15nm for each of the samples analyzed and graph 2 shows the bar graph of normalized area of each peak. In both, it can be noted that there is no pattern of increased intensity of mercury emission due to the increased concentration of the solution deposited in the pellet.

It is possible to observe that although the concentrations were increasing, the peak did not have an increasing behavior for all samples, probably due to the inhomogeneous distribution of the solution in the tablet because its surface was not completely flat, causing the solution to distributed differently in each pellet. Moreover, the concentration spectra that would be used in our experiments do not have a significant intensity to evaluate the peak.

Therefore, as we did not obtain a conclusive result to relate peak intensity to sample concentration and we could not detect the mercury peak in lower concentration samples, we chose not to use the LIBS method to quantify our results.