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Protocols

The protocols used for the experiments performed in the context of our project, are listed below.

1. DNA extraction from soil samples

The Macherey-Nagel™ NucleoSpin™ kit (Fisher Scientific) was used for this experiment.

  1. Sample preparation

    Transfer 500 mg fresh sample material to a MN Bead Tube Type A containing the ceramic beads.

    Add 700 μL Buffer SL1.

  2. Adjust lysis conditions

    Add 150 μL Enhancer SX and close the cap.

  3. Sample lysis

    Attach the MN Bead Tubes horizontally to a vortexer, by taping.

    Vortex the samples at full speed and room temperature (18-25 ℃) for 5 min.

  4. Precipitate contaminants

    Centrifuge for 2 min at 11,000 x g to eliminate the foam caused by the detergent.

    Add 150 μL Buffer SL3 and vortex for 5 s.

    Incubate for 5 min at 0-4 ℃.

    Centrifuge for 1 min at 11,000 x g.

  5. Filter lysate

    Place a NucleoSpin® Inhibitor Removal Column (red ring) in a Collection Tube (2 mL, lid).

    Load up to 700 μL clear supernatant of step 4 onto the filter.

    Centrifuge for 1 min at 11,000 x g.

    Discard the NucleoSpin® Inhibitor Removal Column.

  6. Adjust binding conditions

    Add 250 μL Buffer SB and close the lid.

    Vortex for 5 s.

  7. Bind DNA

    Place a NucleoSpin® Soil Column (green ring) in a Collection Tube (2 mL).

    Load 550 μL sample onto the column.

    Centrifuge for 1 min at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

    Load the remaining sample onto the column.

    Centrifuge for 1 min at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

  8. Wash and dry silica membrane

    1st wash

    Add 500 μL Buffer SB to the NucleoSpin® Soil Column.

    Centrifuge for 30 s at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

    2nd wash

    Add 550 μL Buffer SW1 to the NucleoSpin® Soil Column.

    Centrifuge for 30 s at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

    3rd wash

    Add 650 μL Buffer SW2 to the NucleoSpin® Soil Column.

    Close the lid and vortex for 2 s.

    Centrifuge for 30 s at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

    4th wash

    Add 650 μL Buffer SW2 to the NucleoSpin® Soil Column.

    Close the lid and vortex for 2 s.

    Centrifuge for 30 s at 11,000 x g.

    Discard flow through and place the column back into the collection tube.

  9. Dry silica membrane

    Centrifuge for 2 min at 11,000 x g.

  10. Elute DNA

    Place the NucleoSpin® Soil Column into a new microcentrifuge tube.

    Add 30 μL (for high concentration), 50 μL (for medium concentration and yield), or 100 μL (for high yield) Buffer SE to the column.

    Do not close the lid and incubate for 1 min at room temperature (18-25 ℃).

    Close the lid and centrifuge for 30 s at 11,000 x g.

  11. Quantify DNA

    UV-Vis quantification is performed by using the purity ratios A260/A230 and A260/A280.

    10 μL of each sample were run on a 1 % TAE agarose gel (1 h, 100 V), to verify the UV-Vis quantification.

2. 16S rRNA sequencing

Sequencing was performed using Illumina technology with a MiSeq™PE300 sequencer, following manufacturer's recommendations.

16S rRNA Library Preparation Workflow

  1. Amplicon PCR

    This step uses PCR to amplify templates out of a DNA sample using region of interest specific primers with overhang adapters attached.


    Consumables

    Item Quantity
    Microbial Genomic DNA (5 ng/μl in 10 mM Tris pH 8.5) 2.5 μl per sample
    Amplicon PCR Reverse Primer (1 μM) 5 μl per sample
    Amplicon PCR Forward Primer (1 μM) 5 μl per sample
    2x KAPA HiFi HotStart ReadyMix 12.5 μl per sample
    Microseal 'A' film
    96-well 0.2 ml PCR plate 1 plate

    Procedure

    1. Set up the following reaction of DNA, 2x KAPA HiFi HotStart ReadyMix, and primers:

      Item Quantity
      Microbial Genomic DNA (5 ng/μl in 10 mM Tris pH 8.5) 2.5 μl
      Amplicon PCR Reverse Primer (1 μM) 5 μl
      Amplicon PCR Forward Primer (1 μM) 5 μl
      2x KAPA HiFi HotStart ReadyMix 12.5 μl

    2. Seal plate and perform PCR in a thermal cycler using the following program:

      • 95 ℃ for 3 minutes
      • 25 cycles of:
        1. 95 ℃ for 30 seconds
        2. 55 ℃ for 30 seconds
        3. 72 ℃ for 30 seconds
      • 72 ℃ for 5 minutes
      • Hold at 4 ℃

  2. PCR Clean Up

    This step uses AMPure XP beads to purify the 16S V3 and V4 amplicon away from free primers and primer dimer species.


    Consumables

    Item Quantity
    10 mM Tris pH 8.5 52.5 μl per sample
    AMPure XP beads 20 μl per sample
    Freshly Prepared 80% Ethanol (EtOH) 400 μl per sample
    96-well 0.2 ml PCR plate 1 plate

    Preparation

    Bring the AMPure XP beads to room temperature.


    Procedure

    1. Centrifuge the Amplicon PCR plate at 1,000 × g at 20 ℃ for 1 minute to collect condensation, carefully remove the seal.
    2. Vortex the AMPure XP beads for 30 seconds to make sure that the beads are evenly dispersed. Add an appropriate volume of beads to a trough depending on the number of samples processing.
    3. Using a multichannel pipette, add 20 μl of AMPure XP beads to each well of the Amplicon PCR plate. Change tips between columns.
    4. Gently pipette the entire volume up and down 10 times if using a 96-well PCR plate.
    5. Incubate at room temperature without shaking for 5 minutes.
    6. Place the plate on a magnetic stand for 2 minutes or until the supernatant has cleared.
    7. With the Amplicon PCR plate on the magnetic stand, use a multichannel pipette to remove and discard the supernatant. Change tips between samples.
    8. With the Amplicon PCR plate on the magnetic stand, wash the beads with freshly prepared 80% ethanol as follows:
      1. Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each sample well.
      2. Incubate the plate on the magnetic stand for 30 seconds.
      3. Carefully remove and discard the supernatant.
    9. With the Amplicon PCR plate on the magnetic stand, perform a second ethanol wash as follows:
      1. Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each sample well.
      2. Incubate the plate on the magnetic stand for 30 seconds.
      3. Carefully remove and discard the supernatant.
      4. Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol.
    10. With the Amplicon PCR plate still on the magnetic stand, allow the beads to air-dry for 10 minutes.
    11. Remove the Amplicon PCR plate from the magnetic stand. Using a multichannel pipette, add 52.5 μl of 10 mM Tris pH 8.5 to each well of the Amplicon PCR plate.
    12. Gently pipette, mix up and down 10 times, changing tips after each column (or seal plate and shake at 1800 rpm for 2 minutes). Make sure that beads are fully resuspended.
    13. Incubate at room temperature for 2 minutes.
    14. Place the plate on the magnetic stand for 2 minutes or until the supernatant has cleared.
    15. Using a multichannel pipette, carefully transfer 50 μl of the supernatant from the Amplicon PCR plate to a new 96-well PCR plate. Change tips between samples to avoid cross-contamination.

  3. Index PCR

    This step attaches dual indices and Illumina sequencing adapters using the Nextera XT Index Kit.


    Consumables

    Item Quantity
    2x KAPA HiFi HotStart ReadyMix 25 μl per sample
    Nextera XT Index 1 Primers (N7XX) from the Nextera XT Index kit 5 μl per sample
    Nextera XT Index 2 Primers (S5XX) from the Nextera XT Index kit 5 μl per sample
    PCR Grade Water 10 μl per sample
    TruSeq Index Plate Fixture (FC-130-1005) 1
    96-well 0.2 ml PCR plate 1 plate
    Microseal 'A' film 1

    Procedure

    1. Using a multichannel pipette, transfer 5 μl from each well to a new 96-well plate. The remaining 45 μl is not used in the protocol and can be stored for other uses.
    2. Arrange the Index 1 and 2 primers in a rack, using the following arrangements as needed:
      1. Arrange Index 2 primer tubes (white caps, clear solution) vertically, aligned with rows A through H.
      2. Arrange Index 1 primer tubes (orange caps, yellow solution) horizontally, aligned with columns 1 through 12.
      3. Place the 96-well PCR plate with the 5 μl of resuspended PCR product DNA in the TruSeq Index Plate Fixture.
    3. Set up the following reaction of DNA, Index 1 and 2 primers, 2x KAPA HiFi HotStart ReadyMix, and PCR Grade water:
      4. Volume
      DNA 5 μl
      Nextera XT Index Primer 1 (N7XX) 5 μl
      Nextera XT Index Primer 2 (S5XX) 5 μl
      2x KAPA HiFi HotStart ReadyMix 25 μl
      PCR Grade Water 10 μl
      Total 50 μl
    4. Gently pipette up and down 10 times to mix.
    5. Cover the plate with Microseal 'A'.
    6. Centrifuge the plate at 1,000 × g at 20 ℃ for 1 minute.
    7. Perform PCR on a thermal cycler using the following program:
      • 95 ℃ for 3 minutes
      • 8 cycles of:
        1. 95 ℃ for 30 seconds
        2. 55 ℃ for 30 seconds
        3. 72 ℃ for 30 seconds
      • 72 ℃ for 5 minutes
      • Hold at 4 ℃
    Table 1.Primers and barcode sequences used for library preparation.

  4. PCR Clean Up 2

    This step uses AMPure XP beads to clean up the final library before quantification.


    Consumables

    Item Quantity
    10 mM Tris pH 8.5 27.5 μl per sample
    AMPure XP beads 56 μl per sample
    Freshly Prepared 80% Ethanol (EtOH) 400 μl per sample
    96-well 0.2 ml PCR plate 1 plate

    Procedure

    1. Centrifuge the Index PCR plate at 280 × g at 20 ℃ for 1 minute to collect condensation.
    2. Vortex the AMPure XP beads for 30 seconds to make sure that the beads are evenly dispersed. Add an appropriate volume of beads to a trough.
    3. Using a multichannel pipette, add 56 μl of AMPure XP beads to each well of the Index PCR plate.
    4. Gently pipette mix up and down 10 times if using a 96-well PCR plate
    5. Incubate at room temperature without shaking for 5 minutes.
    6. Place the plate on a magnetic stand for 2 minutes or until the supernatant has cleared.
    7. With the Index PCR plate on the magnetic stand, use a multichannel pipette to remove and discard the supernatant. Change tips between samples.
    8. With the Index PCR plate on the magnetic stand, wash the beads with freshly prepared 80% ethanol as follows:
      1. Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each sample well.
      2. Incubate the plate on the magnetic stand for 30 seconds.
      3. Carefully remove and discard the supernatant.
    9. With the Index PCR plate on the magnetic stand, perform a second ethanol wash as follows:
      1. Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each sample well.
      2. Incubate the plate on the magnetic stand for 30 seconds.
      3. Carefully remove and discard the supernatant.
      4. Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol.
    10. With the Index PCR plate still on the magnetic stand, allow the beads to air-dry for 10 minutes.
    11. Remove the Index PCR plate from the magnetic stand. Using a multichannel pipette, add 27.5 μl of 10 mM Tris pH 8.5 to each well of the Index PCR plate.
    12. If using a 96-well PCR plate, gently pipette, mix up and down 10 times until beads are fully resuspended, changing tips after each column.
    13. Incubate at room temperature for 2 minutes.
    14. Place the plate on the magnetic stand for 2 minutes or until the supernatant has cleared.
    15. Using a multichannel pipette, carefully transfer 25 μl of the supernatant from the Index PCR plate to a new 96-well PCR plate. Change tips between samples to avoid cross contamination.

  5. Library Quantification, Normalization, and Pooling

    Library quantification was performed, using a fluorometric quantification method that uses dsDNA binding dyes.

    The Quant-iT™ PicoGreen™ dsDNA Reagent and Kit was used for this purpose.


    Preparing the DNA standard curve

    1. Prepare a 2 μg/mL stock solution of dsDNA in TE. Determine the DNA concentration on the basis of absorbance at 260 nm (A260) in a cuvette with a 1 cm pathlength; an A260 of 0.04 corresponds to 2 μg/mL dsDNA solution.The lambda DNA standard is diluted 50-fold in TE to make the 2 μg/mL working solution.
    2. For the high-range standard curve from 10 ng/mL to 1 μg/mL, dilute the 2 μg/mL DNA stock solution into microplate wells as shown in Table 2.
    Table 2.Protocol for preparing a high-range standard curve.
    Volume of TE buffer Volume of 2 μg/mL DNA stock Volume of diluted Quant-iT™ PicoGreen™ dsDNA Reagent Final DNA concentration in assay
    0 μL 100 μL 100 μL 1 μg/mL
    90 μL 10 μL 100 μL 100 ng/mL
    99 μL 1 μL 100 μL 10 ng/mL
    100 μL 0 μL 100 μL blank

    For the low-range standard curve from 250 pg/mL to 25 ng/mL, dilute the 2 μg/mL DNA solution 40-fold in TE to yield a 50 ng/mL DNA stock solution and then prepare the dilution series shown in Table 3.

    Table 3.Protocol for preparing a low-range standard curve.
    Volume of TE buffer Volume of 50 ng/mL DNA stock Volume of diluted Quant-iT™ PicoGreen™ dsDNA Reagent Final DNA concentration in assay
    0 μL 100 μL 100 μL 25 ng/mL
    90 μL 10 μL 100 μL 2.5 ng/mL
    99 μL 1 μL 100 μL 250 pg/mL
    100 μL 0 μL 100 μL blank

    Analyzing samples

    1. Dilute the experimental DNA solution in TE to a final volume of 100 μL in microplate wells.
    2. Add 100 μL of the aqueous working solution of the Quant-iT™ PicoGreen™ dsDNA Reagent, which is at room temperature, to each sample. Incubate for 2-5 minutes at room temperature, protected from light.
    3. Measure the fluorescence of the samples using the same instrument parameters used to generate the standard curve.
    4. Subtract the fluorescence value of the reagent blank from that of each of the samples. Determine the DNA concentration of the sample from the standard curve, which was created previously.

    Calculate DNA concentration in nM, based on the size of DNA amplicons as determined by an Agilent Technologies 2100 Bioanalyzer trace:

    Dilute concentrated final library using Resuspension Buffer (RSB) or 10 mM Tris pH 8.5 to 4nM. Aliquot 5 μl of diluted DNA from each library and mix aliquots for pooling libraries with unique indices.

  6. Library Denaturing and MiSeq Sample Loading

    In preparation for cluster generation and sequencing, pooled libraries are denatured with NaOH, diluted with hybridization buffer, and then heat denatured before MiSeq sequencing.


    Consumables

    Item Quantity
    10 mM Tris pH 8.5 or RSB (Resuspension Buffer) 6 μl
    HT1 (Hybridization Buffer) 1540 μl
    0.2 N NaOH (less than a week old) 10 μl
    PhiX Control Kit v3 (FC-110-3001) 4
    MiSeq reagent cartridge 1 cartridge
    1.7 ml microcentrifuge tubes (screw cap recommended) 3 tubes
    2.5 L ice bucket

    Preparation

    1. Set a heat block suitable for 1.7 ml microcentrifuge tubes to 96 ℃.
    2. Remove a MiSeq reagent cartridge from -15 ℃ to -25 ℃ storage and thaw at room temperature.
    3. In an ice bucket, prepare an ice-water bath by combining 3 parts ice and 1 part water.

    Denature DNA

    1. Combine the following volumes of pooled final DNA library and freshly diluted 0.2 N NaOH in a microcentrifuge tube:
      1. 4 nM pooled library (5 μl)
      2. 0.2 N NaOH (5 μl)
    2. Set aside the remaining dilution of 0.2 N NaOH to prepare a PhiX control within the next 12 hours.
    3. Vortex briefly to mix the sample solution, and then centrifuge the sample solution at 280 × g at 20 ℃ for 1 minute.
    4. Incubate for 5 minutes at room temperature to denature the DNA into single strands.
    5. Add the following volume of pre-chilled HT1 to the tube containing denatured DNA:
      1. Denatured DNA (10 μl)
      2. Pre-chilled HT1 (990 μl)
      3. Adding the HT1 results in a 20 pM denatured library in 1 mM NaOH.
    6. Place the denatured DNA on ice until you are ready to proceed to final dilution.

    Dilute Denatured DNA

    1. Dilute the denatured DNA to the desired concentration. We started our first run using 4pM loading concentration.
      2. Final Concentration 2 pM 4 pM 6 pM 8 pM 10 pM
      20 pM denatured library 60 μl 120 μl 180 μl 240 μl 300 μl
      Pre-chilled HT1 540 μl 480 μl 420 μl 360 μl 300 μl
    2. Invert several times to mix and then pulse centrifuge the DNA solution.
    3. Place the denatured and diluted DNA on ice.

    Denature and Dilution of PhiX Control

    Use the following instructions to denature and dilute the 10 nM PhiX library to the same loading concentration as the Amplicon library. The final library mixture must contain at least 5% PhiX.

    1. Combine the following volumes to dilute the PhiX library to 4 nM:
      1. 10 nM PhiX library (2 μl)
      2. 10 mM Tris pH 8.5 (3 μl)
    2. Combine the following volumes of 4 nM PhiX and 0.2 N NaOH in a microcentrifuge tube:
      1. 4 nM PhiX library (5 μl)
      2. 0.2 N NaOH (5 μl)
    3. Vortex briefly to mix the 2 nM PhiX library solution.
    4. Incubate for 5 minutes at room temperature to denature the PhiX library into single strands.
    5. Add the following volumes of pre-chilled HT1 to the tube containing denatured PhiX library to result in a 20 pM PhiX library:
      1. Denatured PhiX library (10 μl)
      2. Pre-chilled HT1 (990 μl)
    6. Dilute the denatured 20 pM PhiX library to the same loading concentration as the Amplicon library.
    7. Invert several times to mix and then pulse centrifuge the DNA solution.
    8. Place the denatured and diluted PhiX on ice.

    Combine Amplicon Library and PhiX Control

    1. Combine the following volumes of denatured PhiX control library and your denatured amplicon library in a microcentrifuge tube:
      1. Denatured and diluted PhiX control (30 μl)
      2. Denatured and diluted amplicon library (570 μl)
    2. Set the combined sample library and PhiX control aside on ice until you are ready to heat denature the mixture immediately before loading it onto the MiSeq v3 reagent cartridge.
    3. Using a heat block, incubate the combined library and PhiX control tube at 96 ℃ for 2 minutes.
    4. After the incubation, invert the tube 1-2 times to mix and immediately place in the ice water bath.
    5. Keep the tube in the ice-water bath for 5 minutes.
    6. Load the library into the MiSeq reagent cartridge.

3. qPCR

Quantitative PCR (qPCR) was performed in a StepOne system (Applied Biosystems; Thermo Fisher Scientific).

The PowerUp™ SYBR™ Green Master Mix (Applied Biosystems™) was used to prepare the PCR samples.

Component Volume (20 μL/well)
PowerUp™ SYBR™ Green Master Mix (2X) 10μL
Forward and reverse primers[1] 5μL
DNA template + Nuclease-Free Water[2] Variable
Total 20 μL

[1] Exact primers used: BACT1369F CGGTGAATACGTTCYCGG and PROK1492R GGWTACCTTGTTACGACTT.

[2] The volume is dependent on the initial concentration of the sample.

Protocol steps:

  1. Swirl the PowerUp™ SYBR™ Green Master Mix to mix thoroughly.
  2. Thaw the DNA samples and primers on ice, vortex to mix, then centrifuge briefly.
  3. Mix the components thoroughly, then centrifuge briefly to spin down the contents and eliminate any air bubbles.
  4. Transfer the appropriate volume of each reaction to each well of an optical plate.
  5. Seal the plate with an optical adhesive cover, then centrifuge briefly to spin down the contents and eliminate any air bubbles.
  6. Place the reaction plate in the real-time PCR instrument.
  7. Set the thermal cycling conditions using the default PCR thermal cycling conditions, specified in the following table.
Table 4.Cycling conditions for qPCR.
Steps Temperature Time
1. UDG activation 50 ℃ 2 min
2. Dual-Lock™ DNA polymerase 95 ℃ 2 min
3. Denaturation 95 ℃ 15 sec
4 Annealing 56 ℃ 15 sec
-Repeat steps [2,3,4] for 40 cycles-
5. Extension 72 ℃ 1 min