DNA extraction and amplicon sequencing

Dna extraction comparison

DNA extractions

Modified from Qiagen DNeasy plant mini kit

Prepare:

1. Heat water micro tube shaker to 65˚C.
2. Get ice.
3. Label:
   1. QIAshredder spin columns
   2. DNeasy mini spin columns
   3. 2 mL Eppendorf’s
   4. 2 mL bead beating tubes
   5. Get tubes that DNA will be eluted in to.

Method:

1. Have pellet of cells in 1.5 ml Eppendorf tube.
2. Add 300 ml 1x buffer TAE.
3. Add quarter PCR tube of beads to bead beating tube (2 ml screw top tube).
4. Transfer all sample to bead beating tube.
5. Use bead beater 2 x 45s and 1 x 30s (setting 6.0 or 30 Hz).
6. Add 400 µl buffer AP1, and 4 µl RNase A.
7. Incubate at 65˚C for 10 mins.
8. Add 130 ml buffer P3. Mix and cool on ice for 5 mins.
9. Centrifuge for 5 mins at 14,000 rpm.
10. Pipette lysate (800 µl) into QIAshredder spin column in 2 ml collection tube. Centrifuge for 2 mins at 14,000 rpm.
11. Transfer flow-through (700 µl) to a new Eppendorf. Add 1.5 volumes buffer AW1 (1050 µl) and mix by pipetting.
12. Add mixture to DNeasy mini spin column in 2 ml collection tube, 700 µl at a time. Centrifuge for 1 min at 4,000 rpm.
13. Discard flow-through. Repeat step 13 if necessary (if volumes are the same as mentioned here, then this will be carried out 3 times).
14. Place spin column into new collection tube. Add 500 ml buffer AW2 and centrifuge for 1 min at 4,000 rpm. Discard the flow-through.
15. Add 500 ml buffer AW2 and centrifuge for 2 mins at 13,000 rpm.
16. Discard flow-through and centrifuge for 1 min at 13,000 rpm.
17. Transfer spin column to Eppendorf.
18. Add 30 ml H₂O for elution. Incubate for 5 mins at room temperature. Centrifuge for 1 min at 13,000 rpm.
19. Optional: Repeat step 19 with buffer AE.
20. Measure DNA on Nanodrop – A₂₆₀/A₂₈₀ ratio should be 1.7-1.9.

Library Prep Modified

PCR 1 – 16S/18S/ITS

All done in class II cabinet, after UV.

1. Dilute stocks of primers to 10 µM (60 µL 100 µM stock primer + 540 µL water)

Use 10 µl 1-2 ng/µl DNA (or 3 µl 5 ng/µl DNA and 7 µl water) → need 10-20 ng DNA.

1. Dilute DNA samples to the same concentration (1-2ng/µl) – measured using Qubit kit.
2. Add 16 µl of master mix to each well of a 96 well PCR plate.
3. Add 10 µl DNA template to each well.
4. PCR cycles.

Primers

PCR cycle conditions

Bead clean-up step

Using AmpliClean Magnetic beads (Nimagen)

1. Bring beads to room temperature and vortex for 30 s to make sure the beads are evenly dispersed.
2. Remove what is needed to an Eppendorf/Universal (4 ml for 96 well plate – 40 µl per PCR reaction).
3. For each 96 well plate prepare 50 ml 80% (“200 proof”) ethanol, made with molecular grade water (40 ml ethanol + 10 ml water – 500 µl per sample).
4. Centrifuge the Amplicon (?) PCR plate at ~1,000 x g at room temperature for 1 min to collect condensation, carefully remove seal.
5. Vortex the beads for 30 s to make sure the beads are evenly dispersed. Tip into sterile reservoir.
6. Add 36 µl (1.4X) of beads to each well of the PCR plate using a multichannel pipette and pipette up and down 10 times to mix. Change tips between samples.
7. Incubate at room temperature for 5 minutes.
8. Place the plate on a magnetic stand for 4 minutes, or until the supernatant has cleared.
9. With the plate on the magnetic stand, use a multichannel pipette (set to 70 µl) to carefully remove and discard the supernatant (try not to scrape the edges of the wells). Change tips between samples.
10. With the plate on the magnetic stand, wash with the freshly prepared 80% ethanol (use a sterile reservoir) as follows:
    1. Using a multichannel pipette, add 200 µl of freshly prepared ethanol to each well.
    2. Incubate the plate on the magnetic stand for 30 s.
    3. Carefully remove and discard the supernatant.
11. Repeat step 10 (ethanol wash).
12. Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol. Look at the plate and remove any remaining ethanol with a single channel pipette. Make sure there is no visible liquid.
13. With the plate still on the magnetic stand, allow the beads to air dry for 10 minutes.
14. Remove the plate from the magnetic stand. Add 40 µl of Qiagen buffer EB (10 mM Tris pH 8.5) to each well of the plate (use a sterile reservoir).
15. Pipette up and down (change tips between rows).
    1. Ensure that there are no bubbles at the bottom of wells! Can pipette up from the bottom and place this on top of the liquid in each well.
16. Incubate at room temperature for 2 mins.
17. Place the plate on the magnetic stand for at least 3 minutes (make sure supernatant has cleared).
18. Using a multichannel pipette, carefully transfer 35 µl of the supernatant to a new 96 well PCR plate.
19. Place the plate on a while piece of paper to check for any bead carry over. Slight carry over (slightly brown) samples will be fine, but if any wells have a lot of carry over then place the contents of that well back into the corresponding well on the plate and wait at least 2 mins before transferring back to the PCR plate.
20. Check 5 µl on a 96 well gel (75 ml, 1%, 10 mins at 100 V). Include some positive control (see Sally) to check for size and quantity.
21. Repeat any failed samples, purify and place in the correct well of the purified plate.

SAFE STOPPING POINT

If you do not immediately proceed to Index PCR, seal plate with adhesive seal and store it at -20˚C for up to a week.

PCR 2 – Index

Arrange the Index 1 and 2 primers in a rack (i.e. the TruSeq Index Plate Fixture) using the following arrangements as needed:

• Arrange Index 2 primer tubes (S5xx) vertically, aligned with rows A through H.
• Arrange Index 1 primer tubes (N7xx) horizontally, aligned with columns 1 through 12.

1. Add 17 µl master mix to each well of a 96 well plate using the multichannel pipette.
2. Add 2.5 µl Index primer 1 (N7xx) to well A-H of the appropriate column.
3. Add 2.5 µl Index primer 2 (S5xx) to well 1-12 of the appropriate row.
4. Add 4 µl PCR product from the previous step to each appropriate well.
5. Gently pipette up and down 10 times to mix (can use mix setting on multichannel pipette).
6. Cover the plate with a foil seal.
7. Centrifuge the plate at ~1,000 x g for 1 min.
8. PCR cycles.
   
   PCR cycle conditions
   95˚C for 3 mins
   
   8 cycles of:
       98˚C for 20 s
       55˚C for 15 s
       72 ˚C for 15 s
   
   72˚C for 5 mins
   Hold at 4˚C (or 12 ˚C if overnight?)
    
9. Centrifuge the Index PCR plate at ~1,000 rpm for 1 min to collect condensation.
10. Check a sub-sample (i.e. A1-A3) 5 µl on a midi gel (1%, 40 mins at 90 V). You could do this with a few samples next to PCR product from before index PCR to show the size shift (i.e. A1 before index, A1 after index, A2 before index, A2 after index).

SequalPrep Normalization Plate (96) Kit

Binding step

1. Add 20 µl SequalPrep Normalization Binding Buffer into the wells of the SequalPrep Normalization plate (using multichannel pipette).
2. Add 20 µl from the PCR plate into the wells of the SequalPrep Normalization plate (using multichannel pipette).
3. Mix by completely pipetting up and down, and briefly centrifuge the plate.
4. Incubate the plate for 1 hour at room temperature to allow binding of DNA to the plate surface.

Washing step

1. Aspirate the liquid from wells (you can transfer to a new PCR plate and freeze as a back-up). Be sure not to scrape the sides of the wells during aspiration.
2. Add 50 µl SequalPrep Normalization Wash Buffer to the wells. Mix by pipetting up and down twice to improve removal of contaminants.
3. Completely aspirate the buffer from wells and discard.

Elution step

1. Add 20 µl SequalPrep Normalization Elution Buffer to each well of the plate.
2. Mix by pipetting up and down 5x and briefly centrifuge the plate. Ensure that the buffer contacts the entire plate coating (up to 20 µl level).
3. Incubate at room temperature for 5 mins.
4. Transfer and pool the purified DNA.
5. Check yield on a Qubit DNA Hs kit using 5 µl. Dilute DNA to 4 nM (using table overleaf).
6. If yield is low, concentrate 100-200 µl using the DNA speed vac (see below) and then re-dilute to 4 nM.
   
   Speed vac – Aim to end up with ~50 µl DNA. Mark on an Eppendorf where 50 µl is. Add appropriate amount of DNA to tube to result in correct concentration in 50 µl. Mark where the liquid reaches. Balance the rotor with opened tubes. Switch to manual, run for 30 mins at 45˚C. Check every 30 mins until the liquid has reached the 50 µl line.
    
7. Store the eluted DNA at -20˚C until further use.
8. Give DNA to genomics lab along with DNA concentration and amplicon size (Final insert size in table below).

Amplicon sizes

Fragment = original PCR product (no adpaters/tags), including primers.

Insert = PCR product + adapters/tags.

Calculation based on Illumina MiSeq library prep protocol, p16.