12S rRNA-Gene Metabarcoding Library Prep: Dual-PCR Method

Ensure that the laboratory is appropriately configured and that staff has appropriate training. See "Guidelines" for more information. Pay attention to the separation of pre and post-PCR spaces and equipment.

Ensure that all reagents are aliquoted in appropriate amounts, and stored according to manufacturers' recommendations. Never pipet directly from reagent stocks.

Prepare the SPRI beads' working solution, and test their efficiency following this protocol.

Serapure Beads Preparation and Testing

Prepare primer working stocks (10μM) for both the first and second PCR steps. Here we use Nextera V2 Kit Sets A, B, C, and D. We advise preparing the indexing primers on 96-well plates according to this configuration:

Indexes_plate.xlsx

We advise adding aliquots of the extracted DNA to a 96-Well PCR plate to facilitate the setup of the PCR reaction. This metadata template will help keep track of the samples, and if indexes are configured as described above, also the identity of sample indexes.

Preparations

Reagents:

(Or equal)

(Or equal)

(Or equal)

(Or equal)

• Custom-designed primers ( Miya et al 2015 ) including: | A | B | C | | --- | --- | --- | | MiFish-U-F_overhang | forward | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGTCGGTAAAACTCGTGCCAGC | | MiFish-U-R_overhang | reverse | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGCATAGTGGGGTATCTAATCCCAGTTTG |

UV for 30 minutes the following:

• 96-well PCR plates (or 8-strip tubes)
• Sharpie
• Pipette tips
• Multichannel pipettes
• Pipettes
• Sterile Nuclease-Free Water

Thaw Platinum, Primers, and nuclease-free water. Keep them in a cooling microcentrifuge tube rack.

PCR reactions are carried out in triplicate 12.5 μl reactions:

A	B
Sterile Nuclease-Free water	3.65
Forward primer (10μM)	0.3
Reverse Primer (10μM)	0.3
Platinum SuperFi	6.25
DNA (1-10 ng)	2
TOTAL	12.5

Seal the 96-well plates and transfer them to thermocyclers.

A	B	C	D
denaturation	94°C	3 minutes
denaturation	94°C	30 seconds
annealing	63°C	30 seconds
extension	68°C	30 seconds
GO TO step 2			39 times
final extension	68°C	10 minutes
HOLD	12°C	HOLD

Pool the three replicates.

Run sample pools on a 2% gel (90V, ~40-60 min) to check the size

of the amplicons and the success of the PCR (5μl/sample).

Preparations

Materials

• Serapure SPRI beads. If not already prepared:
• Magnetic 96-well plate stand
• Anhydrous Ethanol to make a fresh 80% ethanol solution
• Molecular grade water

UV for 30 minutes the following:

• 96-well PCR plates (or 8-strip tubes)
• Sharpie
• Pipette tips
• Multichannel pipettes
• Pipettes
• Sterile Nuclease-Free Water

Remove the magnetic beads from the fridge (allow 30 min to reach room temperature).

Vortex the beads before use.* As you now have about 32.5μl reaction (pooled samples - 5μl used on a gel/Qiaxcel), add 26 μl beads to the product to obtain a ratio of 0.8.

• Pipette up and down about ten times (or until the solution is well mixed – you will see that the color changes).
• Spin tubes down to remove drops from the walls.

Incubate a room temperature without shaking for 5 min. For samples that you know have a low DNA concentration, you can increase this incubation time to 30 min.* Place the plate on the magnetic stand until the supernatant has cleared (~ 3 min).

Remove the supernatant with a multichannel pipette, ensuring to not disturb the beads.

With the samples on the magnetic rack, wash the beads by adding 180 μl of freshly prepared 80% ethanol and incubate for 30s. Carefully remove the supernatant without disturbing the beads.

Repeat the washing step

Remove all residual ethanol using a pipette and air dry, leaving the samples on the magnetic stand (~ 5 min*).

 Remove the plate from the magnetic stand and add 32 μl of nuclease-free water for elution. Gently pipet up and down ten times to resuspend the beads.  Incubate the plate at room temperature for 5 min.

Place the plate back on the magnetic rack for at least 5 min or until the supernatant is cleared. 

Carefully transfer 30 μl of the clear supernatant to a new plate. Seal the plate.

Name the plate: Project, [Gene_name], PCR 1, Post-Purification Plate #, Date,  Initials.

Samples can be stored at -20°C for up to 7 days.

Preparations

Reagents:

• i5 and i7 index plates (10 μM) – If not already prepared: | A | B | C | | --- | --- | --- | | Nextera V2 Index1 | forward | CAAGCAGAAGACGGCATACGAGAT[i7]GTCTCGTGGGCTCGG | | Nextera V2 Index 2 | reverse | AATGATACGGCGACCACCGAGATCTACAC[i5]TCGTCGGCAGCGTC |

UV for 30 minutes the following:

• 96-well PCR plates (or 8-strip tubes)  

• Sharpie 

• Pipette tips 

• Multichannel pipettes  

• Pipettes  

• Sterile Nuclease-Free Water

Thaw Taq, i5 and i7 indexes, and nuclease-free water. Keep them in the  IsoFreeze microcentrifuge tube rack.

Prepare PCR reaction in 25μl reactions:

A	B
Sterile Nuclease-Free water	5
Forward primer (10μM)	2.5
Reverse Primer (10μM)	2.5
2XTaq	12.5
DNA (1-10 ng)	2.5
TOTAL	25

Seal the 96-well plates and transfer them to thermocyclers.

A	B	C	D
denaturation	94°C	3 minutes
denaturation	94°C	30 seconds
annealing	55°C	30 seconds
extension	68°C	30 seconds
GO TO step 2			7X
final extension	68°C	5 minutes
HOLD	12°C	HOLD

Run the product on a 2% agarose gel to check the size of the amplicons and success of the PCR (5μl).

Preparations

Materials

• Prepared serapure SPRI beads.
• Magnetic 96-well plate stand
• Anhydrous Ethanol to make a fresh 80% ethanol solution
• Molecular grade water

UV for 30 minutes the following:

• 96-well PCR plates (or 8-strip tubes)
• Sharpie
• Pipette tips
• Multichannel pipettes
• Pipettes
• Sterile Nuclease-Free Water

Remove the magnetic beads from the fridge (allow 30 min to reach room temperature).

Vortex the beads before use.* Add 16 μl beads to 20 μl of PCR product to obtain a ratio of 0.8.

• Pipette up and down ten times (or until the solution is well mixed – you will see that the color changes).
• Spin tubes down to remove drops from the walls.

Incubate a room temperature without shaking for 5 min. For samples that you know have a low DNA concentration, you can increase this incubation time to 30 min.* Place the plate on the magnetic stand until the supernatant has cleared (~ 3 min).

Remove the supernatant with a multichannel pipette, making sure not to disturb

the beads.

With the samples on the magnetic rack, wash the beads by adding 180 μl of freshly prepared 80% ethanol.* Incubate for 30 s.

• Carefully remove the supernatant without disturbing the beads.

Repeat the washing step.

Remove all residual ethanol using a pipette and air dry, leaving the samples on the magnetic stand (~ 5 min*). Keep an eye on the beads and do not over-dry, otherwise, you will not get an efficient DNA recovery.

Remove the plate from the magnetic stand and add 28 μl of nuclease-free water for elution.* Gently pipette up and down ten times to resuspend the beads.

• Incubate the plate at room temperature for 5 min.

Place the plate back on the magnetic rack for at least 5 min or until the supernatant has cleared.

Carefully transfer 25 μl of the clear supernatant to a new plate. Seal the plate.

Name the plate: Project, [Gene_name], PCR 1, Post-Purification Plate #, Date,  Initials.

Samples can be stored at -20°C for up to 7 days.

Use a fluorometric quantification method that uses dsDNA dyes to measure the concentration of your libraries (Qubit or plate reader). If using Qubit, give preference to the broad range kit if you visualize a strong band in the gel: OR

Citation

Samples will have approximately similar concentrations (usually). Re-check samples that showed very high or low concentrations on Qubit/plate reader quantification. 

Calculate sample volume to have a final amount of 10-40 ng. This amount may vary depending on the overall quantification. For example, if on average the concentration of your samples is about 3 ng/μl and you have 20 μl of product,  you can calculate the volume to make up to 60 ng per sample.

Measure the final library pool concentration on Qubit using

Label tube: [Gene_name], [Project_Name], Pooled Amplicons. Date, Initials, pool concentration.

Materials

• Magnetic beads* (SPRI beads or AMpure XP, find aliquots in fridge)

• Magnetic rack for 1.5-2 mL tubes

• Anhydrous Ethanol to make a fresh 80% ethanol solution

• Sterile Nuclease-Free water UV for 30 minutes the following:

• 1.5 mL lo-Bind tubes

• Sharpie

• Pipette tips

• Pipettes

• Sterile Nuclease-Free Water

Remove the magnetic beads from the fridge (allow 30 min to reach room temperature).

Aliquot your pool into 1.5 mL tubes (try to keep similar volumes between aliquots, a maximum of 350 μl per tube). If you have a 2 mL pool, you can aliquot 350 μl pool in 5 tubes, and have the 6th tube with 250 μl.

Calculate the volume of beads to add into each tube (0.8x beads). For the tubes with 350 μl – add 280 μl beads, and for the tube with 250 μl, add 200 μl beads. You will obtain a ratio of 0.8. Pipette up and down ten times (or until the solution is well mixed – you will see that the color changes). Spin tubes down to remove drops from the walls. Incubate a room temperature without shaking for 5 min.

Place the tubes on the magnetic rack until the supernatant has cleared (~ 3 min).

Remove the supernatant with a P1000 pipette, making sure not to disturb the beads.

With the samples on the magnetic rack, wash the beads by adding 500 μl of freshly prepared 80% ethanol and incubate for 30 s. Carefully remove the supernatant without disturbing the beads.

Repeat washing step

Remove all residual ethanol using a pipette and air dry, leaving the samples on the magnetic rack (~ 5 min*). Keep an eye on the beads and do not over dry, otherwise you will not get an efficient DNA recovery.

Remove the tubes from the magnetic rack and add 40 μl of nuclease-free water for elution. Gently pipette up and down ten times to resuspend the beads. Incubate the tubes at room temperature for 5 min.

Place the tubes back on the magnetic rack at least 5 min or until the supernatant is cleared.

Carefully transfer 30 μl of the clear supernatant to a single new tube (pooling the volumes of the 6 tubes).

Repeat the bead cleanup to reduce the volume even more. You may now have 180 μl in your pool, so you must add 144 μl of beads to have a rate of 0.8.

Make final elution in 50 μl of water, transferring 45 μl of the clear supernatant to a new tube.

Name the tube: [Gene_name], Project, Concentrated Pool, Date, Initials.

Materials:

(or similar)

Prepare 1.5-2 L of 1x TBE buffer and autoclave it. Wait for it cool down to room

temperature. *safe stopping point.

Prepare a 3% agarose gel. It may polymerize faster that the gels that you are used to prepare. You may need about 120 mL of buffer and 3.6 g of agarose if using the small electrophoresis system. Add 6 μl of RedSafe. Use a comb with large teeth – it may accommodate a higher volume of product.

If you have 45 μl of product, add 8 μl of loading dye to the tube and mix well. Load the entire tube’s content in the well. You may need more than one well to load the entire content.

Load the Ladder 100 bp in the first and last well (~3 μl).

Run the gel for about 1h at 90V. In this mean time, weigh one 1.5 mL Eppendorf tube and record the weight. Place on your counter: sterile blade for scalpel, a scalpel, and the LED transilluminator.

When the gel run is done, transfer the gel to the transilluminator, and excise the 350 bp band. Place it in the 1.5 mL tube that you have weighed. * Sometimes the stronger band is the non-target one. That’s why it is important to have the ladder running beside your samples to help you to find the right band. See Figure 1.

Weigh the tube with the gel slice. Record the weight.

Use the Promega Wizard SV gel and PCR clean-up to perform the gel purification (follow instructions in the manual - https://www.promega.ca/products/nucleic-acid-extraction/clean-up-and- concentration/wizard-sv-gel-and-pcr-clean-up-system/?catNum=A9281#protocols).

Add 10 μl Membrane Binding Solution per 10 mg of gel slice (for example, if you weight 102 mg of gel slice in the tube, add 102 μl of Membrane Binding Solution).

Vortex and incubate at 50-65oC until gel slice is completely dissolved (~10 min). Vortex again to be sure that you do not have any gel in the tube. Spin down.

Insert SV minicolumn into a collection tube.

Transfer dissolved gel mixture to the minicolumn assembly. Incubate at room temperature for 1 min.

Centrifuge at 16,000 x g for 1 min. Discard flowthrough, and reinsert the minicolumn into thecollection tube. *dry the collection tube edges using Kim wipes if necessary.

Add 700 μl Membrane Wash Solution (ethanol added). Centrifuge at 16,000 x g for 1 min. Discard flowthrough, and reinsert the minicolumn into collection tube.

Repeat the step 15 with 500 μl Membrane Wash Solution. Centrifuge at 16,000 x g for 5 min.

Empty the collection tube and recentrifuged the column assembly for 1 min to allow evaporation of any residual ethanol.

Carefully transfer minicolumn to a clean 1.5 ml tube.

Add 35-50 μl of elution buffer (or Nuclease-Free water) to the minicolumn. Incubate at room temperature for 1 min. Centrifuge at 16,000 x g for 1 min. Perform a second elution if you consider it to be necessary.

Quantify the purified product on Qubit using the ds DNA BR kit.

Label tube: [Gene_name], [Project_Name], Final Library. Date, Initials, concentration.

Library fragment size (BP) is determined using

Molarity of the final pool is assessed using

COI libraries are sequenced an a MiSeq instrument using:

with pair-end setup (2*300 bp), spiked with 10% .