TITLE: Metabarcoding Fecal Swabs or Stomach Contents for Fish and Crustaceans using 2-PCR protocol and Illumina MiSeq
AUTHORS: Eldridge Wisely

[DESCRIPTION]
This protocol describes a method to metabarcode a 170bp region of the mitochondrial16S rRNA gene of crustaceans and a 163-185bp region of the mitochondrial 12S rRNA gene of fishes.  These regions are subjected to PCR separately in multiple replicates and the resulting PCR products are pooled by sample and then indexed for sequencing on an Illumina MiSeq platform.

[BEFORE_START]
Work in a pre-PCR lab, as separated as possible from post-PCR products.

Clean work area with 10% bleach solution before beginning work for the day, then change gloves so that no bleach carryover to your samples or reactions occurs.

[GUIDELINES]
The PCR conditions described here are different from the PCR conditions described by Miya et al., and Berry et al. in their respective publications introducing the primers used here.  This difference is due to the use of the Takara High Fidelity PCR EcoDry Premix in this protocol.

[STEPS]
SECTION: Prepare Primers
1. Order metabarcoding primers with diversity spacers and Illumina overhang sequences (Illumina, 2013):  (Miya et al., 2015):  
TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGNNNNGTCGGTAAAACTCGTGCCAGC
 
  (Miya et al., 2015): GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGNNNNCATAGTGGGGTATCTAATCCCAGTTTG

  Berry et al., 2017): TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGNNNGGGACGATAAGACCCTATA

    (Berry et al., 2017): GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGNNNATTACGCTGTTATCCCTAAAG

We got ours from https://www.idtdna.com/ as 4 nmole Ultramer DNA Oligos, with standard desalting.



SECTION: Prepare Primers
2. Briefly centrifuge primer tubes, then reconstitute primers to 100 micromolar (µM) stock solutions by adding 40 µL  (number of nano moles of primer *10) of  

SECTION: Prepare Primers
3. Make 5 micromolar (µM) working solutions of each primer by adding 95 µL of   and 5 µL of primer stock solution for each 100 µL of primer that you intend to use within the next week or so.  



SECTION: MiFish Takara PCR Recipe
5. Make your MiFish Mastermix:


For each PCR replicate of each sample you intend to process (+10% overage), mix:
0.7 µL   5 micromolar (µM)    
0.7 µL   5 micromolar (µM)   
22.6 µL   





For a full plate of 96 reactions, multiply 105.6*the per-sample volumes in the recipe to make the mastermix.

SECTION: MiFish Takara PCR Recipe
5.2. Add 1 µL DNA extracted from stomach contents or fecal swabs.
SECTION: Create Plate Map
4.1. Do not mix sample types between invasively sampled methods (fecal swabs, or stomach contents) and non-invasively sampled methods (eDNA from water or sediment) in the same PCR procedure.  And don't plan to sequence both types in the same sequencing run with the combinatorial indexing scheme used here.  The potential for contamination of the lower quantity eDNA samples by the higher quantity fDNA samples is too high.  
SECTION: Create Plate Map
4. Determine which sample will go into each well.  This should be the same for each primer set and each replicate.  Include at least one extraction control (you can combine aliquots of the extraction controls from each round of DNA extraction into one tube, and use that as your single extraction control), and include a PCR negative control for each plate of PCR.  See example below of 21 samples, a field negative sample, a combined extraction control, and a PCR negative.
 

SECTION: MiFish Takara PCR Recipe
5.4. Cap each row of reaction tightly before beginning any other PCR reaction in the same room.
SECTION: Crustacean_16S Takara PCR Recipe
7. Make your Crustacean_16S Mastermix:

For each PCR replicate of each sample you intend to process (+10% overage), mix:
2 µL 5 micromolar (µM)   
2 µL 5 micromolar (µM)   
20 µL   



For a full plate of 96 reactions, multiply 105.6*the per-sample volumes in the recipe to make the mastermix.


SECTION: MiFish Takara PCR Recipe
5.3. Mix and stir together with pipette tip, swirling to make sure the liquid is in the bottom, and bringing any bubbles to the surface of each reaction.
SECTION: Crustacean_16S Takara PCR Recipe
8. Add 24 µL  of your Crustacean_16S metabarcoding mastermix to each well of   
SECTION: Crustacean_16S Takara PCR Recipe
8.1. Add 1 µL DNA extract


SECTION: Crustacean_16S Takara PCR Recipe
8.2. Mix and stir together with pipette tip, swirling to make sure the liquid is in the bottom, and bringing any bubbles to the surface of each reaction.
SECTION: Crustacean_16S Takara PCR Recipe
8.3. Cap each row of reaction tightly before beginning any other PCR reaction in the same room.
SECTION: MiFish Takara PCR Conditions
6. 95 °C for 1 min 
35 cycles of:
95 °C for 30 s 
66 °C for1 min 
followed by:
68 °C for1 min 
Hold at 4 °C 


SECTION: Crustacean_16S Takara PCR Conditions
9. 95 °C for 1 min 
35 cycles of:
95 °C for 30 s 
50 °C for 1 min 
68 °C for 30 s 
followed by:
68 °C for 1 min 
then hold at 4 °C 

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
16. In the bead-cleanup plate, do the following steps for one 8-sample row of the plate at a time, pulling back the foil cover for each row after the previous one has been completed.

SECTION: Prepare EtOH for bead cleanup, and bring beads to room temperature
14. Make fresh 80% EtOH so that you will have at least 200 µL of EtOH per well of the combined plate.

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
17. After the 5 min incubation, place 96-well plate on a 
 
 for2 min or until liquid is clear.

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18. remove and discard liquid from the row, being careful not to touch the beads with the pipette or to let the beads dry for more than 30 seconds.

Tip: If you do get beads in your pipette tip, just put the liquid and beads back into the well and wait until the solution clears before trying again.
SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.1. Add 100 µL of fresh 80%EtOH to each well of beads, without disturbing the beads or removing the plate from the magnet. Incubate at Room temperature for 30 s


SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.2. Remove the EtOH, then immediately add another 100 µL of 80% EtOH to the wells, incubate for 30 s Room temperature .

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.3. Remove ALL EtOH, and let the row of beads dry just enough to lose some shine but not enough to start cracking.  This should be approximately 30 s to 1 min.

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
16.1. Add 1.5x the sample volume of Ampure XP beads.  

In this case you have 60uL of combined PCR products per well, so you will add 90 µL of room temperature, well-mixed   to each well, and pipette mix very thoroughly, by stirring and pipetting up and down ten times.

Incubate 5 min  at room temperature.
SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.4.  Remove the plate with new row of cleaned beads from the magnetic plate, and add 30 µL of  
to each well of beads, pipette mixing each well thoroughly.  Incubate 5 min at Room temperature

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.6. Roll back the foil on the final cleaned reactions plate for the appropriate row.  Remove  28 µL clear eluate from the bead-cleanup plate, and place in the appropriate wells of the final cleaned reactions plate.  Immediately cover this cleaned PCR product with 8-strip caps.

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.5. Place back on the magnetic rack for 1 min until liquid is clear again.

SECTION: Perform a 1.5x bead cleanup with Ampure XP beads.
18.7. uncover the next row of samples for cleaning and   
 


SECTION: Prepare Indexing PCR
20. Create an indexing plate map and make sure your chosen indexes (iNext indexes) are color balanced if you aren't doing full 96-well plates at one time.

 

See: Glenn, Travis C., Roger A. Nilsen, Troy J. Kieran, Jon G. Sanders, Natalia J. Bayona-Vásquez, John W. Finger, Todd W. Pierson, et al. 2019. “Adapterama I: Universal Stubs and Primers for 384 Unique Dual-Indexed or 147,456 Combinatorially-Indexed Illumina Libraries (iTru & iNext).” PeerJ 2019 (10). https://doi.org/10.7717/peerj.7755. Supplemental file S10
for information and spreadsheet to help color-balancing the indexes.


SECTION: Gel to check Indexing PCR Products
26. Visualize PCR products in a 1.7-2% gel.  Bands should be around 350-400bp.
SECTION: Prepare Indexing PCR
23. In a new, clean 96-well plate (UV before use if possible and prepare in a pre-PCR space):
 
Add 8.1 µL Indexing Mastermix to each well that will be used and add 0.7 µL of the 5 micromolar (µM) iNext forward indexed primer for each horizontal row of the plate (8 letters), and 0.7 µL 5 micromolar (µM) of the iNext reverse indexed primer for each vertical column of the plate (12 numbers) according to the indexing plate map.
SECTION: Prepare Indexing PCR
22. Indexing PCR Mastermix Recipe:
6 µL   
2.1 µL  
 per sample.  

Multiply by number of wells *10% as explained above, to create master mix.
SECTION: Prepare Indexing PCR
24. Take the prepared indexing reactions to the post-PCR space to add the cleaned PCR product.

In the post-PCR area, add 2.5uL of cleaned PCR 1 product to their associated wells from the indexing plate map.
SECTION: Indexing PCR Conditions
25. 95 °C 3 min
8 cycles of:
98 °C 20 s
65 °C 15 s
72 °C 15 s
final extension of:
72 °C1 min

then hold 4 °C 


SECTION: Combine and Clean all indexed samples from each plate
27. Combine 10uL of up to 70 indexed samples (library) into a single 1.5mL (or 1.7mL) tube.  If there are more than 70 samples, you will need another tube.  
SECTION: Combine and Clean all indexed samples from each plate
28. Multiply the volume of the pooled libraries in each tube by 0.9 to get the volume of Ampure XP beads needed to clean up the reactions.  

For 70 uL of combined libraries you will need 63uL of beads for a total of 133uL of beads+library pool.
SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
29. In the 1.5mL tube of pooled libraries, add 0.9x volume of Ampure XP beads and pipette mix well.  incubate Room temperature for 10 min

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
30. Make enough fresh 80% EtOH to have 2x the total volume of the beads+library pool plus a bit extra.
SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
31. Place 1.5mL tube into a magnetic rack
 
 and incubate Room temperature for 5 min

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
32. Discard liquid and add an equal or greater volume of 80% EtOH.  Incubate Room temperature for 1 min

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
33. Repeat the ethanol wash a second time   , then after the second 80% EtOH wash, remove all EtOH and dry the beads slightly (just until no longer wet-looking but not cracking either).

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
34. Resuspend beads with 100 µL   by pipette mixing thoroughly.  Incubate Room temperature10 min

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
35. Place 1.5 mL tube back on magnet rack and wait until liquid is clear, approximately 2 min

SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
36. remove 100uL of the clear eluate from the tube with beads while on the magnet and place in a new 1.5mL tube.
SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
38. Quantify with Qubit Broad range and visualize in a gel, then send an aliquot of the cleaned, pooled libraries for sequencing.  Check with the sequencing core you're working with for their minimum concentration and volume and try to exceed it by a decent (10-50%) margin if possible.  
SECTION: Prepare EtOH for bead cleanup, and bring beads to room temperature
13. Get AmpureXP beads out of the refrigerator, and bring to room temp, swirl to mix occasionally, or use a rocking platform.
SECTION: Visualize PCR Products
12. Make a 1.7% to 2% agarose gel and run a representative sample of reactions on it to make sure the PCRs worked, producing bands in the 250-300bp range.  Use a ladder than allows you to distinguish small bands between 100-500bp.  Check some PCR negatives to see that they don't have bands.  Be very careful opening the PCR plate wells at this point to avoid cross-contamination.
SECTION: Prepare EtOH for bead cleanup, and bring beads to room temperature
15. Get 2 sterile DNAase/RNAse free 96-well PCR plates out of their packaging and immediately cover with adhesive foil or if possible, UV clean the plates for 15 min, then immediately cover with adhesive foil.

One plate will be for the bead-cleanup steps, and the other will be for the final, cleaned reactions.
SECTION: Combine PCR Products of Biological Samples by Sample and Negatives by Primer
10. Briefly vortex to mix and lightly centrifuge PCR products to get any bubbles from the bottom and any droplets off of the lids.
8-strip tubes can be put into a plate holder and spun down as a plate if this option is available.  
SECTION: Combine PCR Products of Biological Samples by Sample and Negatives by Primer
11. Get a new sterile 96-well plate out of its packaging and immediately cover with foil (a 15-minute treatment under a UV light is helpful to sterilize before covering with foil).
SECTION: Combine PCR Products of Biological Samples by Sample and Negatives by Primer
11.1. For biological samples (not PCR negative controls): combine 10 µL of each of the 6 PCR products by sample into the new sterilized plate.  
 
Peel back one row of foil at a time and using a multitip pipette, open only one row of PCR products at a time to combine.  Cap the resulting combined row of the new plate with a strip-cap and close the PCR products of the previous row before opening a new row.

This should result in 60uL of an equal volume of MiFish and Crustacean-16S PCR products, each from the same original sample.
SECTION: Combine PCR Products of Biological Samples by Sample and Negatives by Primer
11.2. For negative controls: combine 20 µL of each of the 3 negative PCR controls by primer.  You should have one MiFish combined negative and one Crustacean-16S negative, each with a total of 60 µL

SECTION: Visualize PCR Products
12.1. Run the gel at 100V until the dye band has traveled ¾ of the length of the gel, then visualize.
SECTION: Visualize PCR Products
12.2. Any DNA smaller than 250 bp will be excess primer and oligos and will need to be cleaned with SPRI beads.  Check that the initial PCR worked before continuing on with the library-building procedure. 
SECTION: Perform a 0.9x bead cleanup with Ampure XP beads
37. If you had more than one 1.5mL tube, combine equal volumes of the resulting cleaned pooled libraries (the clear eluates) into a new tube.
SECTION: Send for Illumina MiSeq Sequencing
40. Send 100uL of cleaned, pooled libraries for sequencing on the MiSeq platform.  (This combinatorial indexing strategy is not compatible with NovaSeq).  

Tip: We have sequenced up to 1.75 plates of diet samples per MiSeq run, and still gotten sufficient coverage.
SECTION: Optional Quantification and Visualization
39. If you have a Qubit available, quantify with the Broad Range chemistry so that you know that the aliquot you send for sequencing is at least the sequencing core's minimum concentration.  If it's below that concentration,    and elute in 30 µL instead of 100uL in step 34.

SECTION: MiFish Takara PCR Recipe
5.1. Add 24 µL  of your MiFish metabarcoding mastermix to each well of   
SECTION: Prepare Indexing PCR
19. Work in a pre-PCR area, preferably a cleaned and UV-sterilized hood to prepare your indexing reactions before going into the post-PCR area to add the PCR1 products.  
SECTION: Prepare Indexing PCR
21. Prepare working solutions of 5 micromolar (µM) of each indexing primer you intend to use.