ddRADseq Library Preparation Cheat Sheet
Written by Steve Doyle (s.doyle@latrobe.edu.au) - Feb 2014
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simplified version of ddRADseq from Peterson et al 2012 PLoS One
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it is a long protocol, so make sure you read and understand it before starting
o make note of incubation times and safe stopping points to ensure you give
yourself enough time to complete the protocol
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keep track of regents used, either when completing complete protocol or optimizing
particular steps
Sample Setup for up to 96 wells
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it is good practice to make a plate map with sample names clearly shown to ensure
there is no confusion, particularly during barcoding/indexing steps
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suggest you setup an excel document to keep track of samples and barcodes, as well
as data collected along the way, eg. DNA concentrations. It also simplifies calculating
sample volumes for pooling etc. I have a template for this if wanted.
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STEP 1: DNA digestion
- see appendix notes on sample preparation
Preparation
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thaw on ice
o cutsmart buffer
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put restriction enzymes on ice
1. label a 96-well round bottom plate
2. add DNA to wells, adjusting final volume to 20 ul with H2O
3. prepare the following restriction digest master-mix
Reagent
Volume (1x)
Master-mix Volume
(# samples +1 extra)
Cutsmart Buffer
3
EcoRI HF
1
MspI
1
H2O
5
4. Aliquot 10 ul of restriction digest master-mix to each well, pipetting up and down
10X to mix
5. Cover plate with a plate seal
6. Vortex plate for 10 seconds
7. centrifuge at 280g at 20C for 1 min
8. incubate at 37C for at least 6 hours (overnight is fine)
a. we have routinely digested for 3-4h, however, the MspI sometimes needs
longer than this to finish digestion
STEP 2 : Ligation of adapters (starting volume: 30 ul)
Preparation
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thaw on ice
o T4 DNA ligase buffer (note: this is sensitive to freeze/thaw cycles – it is best
that this is aliquoted)
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put T4 DNA ligase on ice
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remove P1 adapter plate from fridge/freezer and allow to come to room
temperature
o vortex and pulse spin plate before opening
1. prepare the following ligation master-mix
Reagent
Volume (1x)
Master-mix Volume
(# samples +1 extra)
T4 DNA ligase buffer
4
T4 DNA ligase
1
Adapter P2 (2 uM stock)
2
H2O
1
2. aliquot 8 ul of ligation master-mix to each well and pipette up and down 10X to mix
3. add 2 ul of each UNIQUE Adapter P1 (2 uM stock) to each well using multichannel
pipette and pipette up and down to mix
4. Cover plate with a plate seal
5. Vortex plate for 10 seconds
6. centrifuge at 280g at 20C for 1 min
7. incubate at 16-20C for 1 hour
a. 16C incubator is ideal, room temperature is ok
8. add 2 ul of 0.5 M EDTA to stop ligation
9. ***SAFE STOPPING POINT – PUT IN FRIDGE FOR UP TO 2 DAYS ***
STEP 3: Library clean & bead double size selection (0.5X + 0.70X size
selection; starting volume: 42 ul in plate)
- see appendix notes on double size selection
Preparation
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make PEG buffer (20% PEG w/v, 2.5M NaCl)
o 10 g PEG 6000
o 7.305 g NaCl
o top up to 50 ml with MilliQ H2O
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make Ampure mix 1
Reagent
Volume (1x)
Master-mix Volume
(# samples +4 extra)
Ampure XP
10
PEG buffer
40
- make Ampure mix 2
Reagent
Volume (1x)
Master-mix Volume
(# samples +4 extra)
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Ampure XP
10
PEG buffer
10
Prepare fresh Ethanol (70% if AmpureXP protocol, 80% if Illumina protocol) – 400 ul
per sample
1. add 58 ul H2O to each well to bring the total volume per well to 100 ul
2. add 50 ul of ampure mix 1 to each well and pipette up and down 10X to mix
3. incubate at least 30 mins on bench
4. place plate on magnet for 5 mins (make sure liquid appears clear)
5. in A NEW PLATE (label PLATE2), add 20 ul of ampure mix 2 to each well ready for
sample
6. while the original plate is on the magnet, transfer the SUPERNATANT to the new
plate OFF the magnet containing the 20 ul ampure mix 2, and pipette up and down
10X to mix
7. Incubate off the magnet for at least 30 mins
8. Transfer plate to magnet and Incubate for 5 mins (at least until solution clears)
9. Discard supernatant by pipette / aspirate
10. Wash samples by adding 190 ul of 70-80% Ethanol to each sample and wait 30
seconds
11. Discard Ethanol by pipette / aspirate
12. Repeat steps 8 & 9 for total of two washes
13. Remove from plate from magnet and allow to air dry for 2-3 mins – make sure you
cannot detect any ethanol
a. NOTE: do not allow plate to dry for too long – beads will start to crack and
become hard to resuspend
b. Not a huge problem if this happens, but makes it more difficult as mixing
steps following become more difficult – keep going though
14. Once ethanol has evaporated, add 12 ul of H20 and pipette up and down 10X to mix,
making sure beads have resuspended
15. allow to incubate for at least 10 mins
16. Place plate back on magnet for 5 mins (at least until solution turns clear)
17. Transfer supernatant containing DNA to a new plate for storage of UNAMPLIFIED
library
STEP 4: PCR amplification of libraries
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see appendix for notes on PCR
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Use the plate map below to design the indexing strategy, and ensure that for
samples that have the same barcode, they have a different index
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1. setup PCR master-mixes, making sure they differ in P2 primer if needed
Reagent
Volume (1x)
Master-mix Volume
(# samples +1 extra)
KAPA HiFi Real Time master
10
mix
PCR1 P1 primer (10 uM)
0.5
PCR2 P2 primer (10 uM)
0.5
2. add 11 ul of PCR master-mix to each well of a 96-well PCR plate
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3. add 9 ul of DNA from the UNAMPLIFIED plate to corresponding well of PCR plate
4. Cover plate with a plate seal (NOTE: must be optically clear seal for qPCR)
5. Vortex plate for 10 seconds
6. centrifuge at 280g at 20C for 1 min
7. qPCR machine setup (ask if you have not been shown how to use it before)
a. PCR protocol
i. 98C – 2 mins
ii. 20 cycles of
1. 98C - 15 secs
2. 60C – 30 secs
3. 72C – 30 secs
b. IMPORTANT: even though you have set the protocol for 20 cycles, you are
not necessarily going to run it for this long
i. You need to watch the qPCR reaction and wait for the amplification of
products
ii. When all samples start to show amplification (approaching
exponential part of amplification curve), STOP THE RUN!
iii. Do not let samples reach plateau
iv. Ideally, this will be between 10-15 cycles, but may need 18 cycles
8. ***SAFE STOPPING POINT – PUT IN FREEZER ***
STEP 5: Purification of PCR-amplified libraries
Preparation
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make Ampure mix
Reagent
Volume (1x)
Master-mix Volume
(# samples + 4 extra)
Ampure XP
10
PEG buffer
60
- Prepare fresh Ethanol (70% if AmpureXP protocol, 80% if Illumina protocol) – 400 ul
per sample
1. add 70 ul of ampure mix to new 96-well round bottom plate
2. add 80 ul of H2O to each well of the PCR plate to bring the total volume per well to
100 ul
3. transfer 100 ul of sample to the 96-well plate containing 70 ul ampure mix and
pipette up and down 10X to mix
4. incubate at least 30 mins on bench
5. place plate on magnet for 5 mins (make sure liquid appears clear)
6. Discard supernatant by pipette / aspirate
7. Wash samples by adding 190 ul of 70-80% Ethanol to each sample and wait 30
seconds
8. Discard Ethanol by pipette / aspirate
9. Repeat steps 8 & 9 for total of two washes
10. Remove from plate from magnet and allow to air dry for 2-3 mins – make sure you
cannot detect any ethanol
a. NOTE: do not allow plate to dry for too long – beads will start to crack and
become hard to resuspend
b. Not a huge problem if this happens, but makes it more difficult as mixing
steps following become more difficult – keep going though
11. Once ethanol has evaporated, add 20 ul of H20 and pipette up and down 10X to mix,
making sure beads have resuspended
12. allow to incubate for at least 10 mins
13. Place plate back on magnet for 5 mins (at least until solution turns clear)
14. Transfer supernatant containing DNA to a new plate labelled “AMPLIFIED library” for
storage
STEP 6: Preparation for, and pooling of samples (starting volume =
20 ul)
- see appendix notes on pooling libraries
1. qubit all samples (record concentrations below)
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2. determine a suitable amount of DNA per sample to pool, keeping in mind to
maximize the amount of DNA per sample as possible
3. Pool samples in a single microcentrifuge tube and determine the volume of the total
pool
4. If volume is above 1000 ul, split into multiple microcentrifuge tubes so that the total
volume per tube is not greater than 500 ul
5. If less than 500ul, top up with H2O so that the total volume is 500 ul
6. Add 50 ul Ampure XP beads and 300 ul PEG buffer to the pooled tube(s)
7. Pipette up and down 10X to mix and allow to incubate for at least 30 mins
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8. Place microcentrifuge tube in tube magnet and incubate for 5 mins (at least until
solution clears)
a. If you had to split the initial pooled DNA into multiple tubes, you will need to
make an additional step.
b. After the 5 minute incubation, discard the supernatant from each tube, and
remover from the magnet
c. Resuspend beads in each tube using 100 ul PEG6000 buffer, and then
combine all bead supernatants into a single tube
d. Place back on the tube magnet for 5 minutes (at least until solution clears)
9. Discard supernatant
10. Wash samples by adding 500 ul of 70-80% Ethanol to each sample and wait 30
seconds
11. Discard Ethanol by pipette / aspirate
12. Repeat steps 8 & 9 for total of two washes
13. Remove from plate from magnet and allow to air dry for 2-3 mins – make sure you
cannot detect any ethanol
a. NOTE: do not allow plate to dry for too long – beads will start to crack and
become hard to resuspend
b. Not a huge problem if this happens, but makes it more difficult as mixing
steps following become more difficult – keep going though
14. Once ethanol has evaporated, add 150 ul of H20
a. Pipette up and down 20X to mix and allow to incubate for at least 10 mins
15. Place plate back on magnet for 5 mins (at least until solution turns clear)
16. Transfer supernatant containing DNA to a new microcentrifuge tube labelled
POOLED AMPLIFIED LIBRARY
STEP 7: Size selection of libraries
- see appendix notes on size selection of libraries
Preparation
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make a 1.5% agarose gel
o use tape on the comb to join 3-4 wells together to make a “super-well” (see
appendix image for example)
o allow for ladder loading wells on both sides of the super-well, with a spare
well in-between ladder and library wells
o make gel as per usual – we use GelRed as a nucleic acid stain – SYBR green or
others will be fine.
1. Add sufficient loading dye to 150 ul of library prior to sample loading
2. once gel is made, load libraries into super-wells. You might need to split library
across two super-wells
a. be conscious of DNA concentration of pooled library – I would not load more
than 1 ug of DNA per super-well as it might artificially smear
3. load 100-bp ladder on both sides of library
4. run gel at 80V for 100 mins
5. check that library has run sufficiently – be quick as you do not want to over expose
DNA to UV light – run gel for longer if needed
6. turn on UV light and QUICKLY mark gel with gel cutting instrument (blade, coverslip,
gel cutting tool) to indicate the size selection, and turn off UV light
7. cut gel fully, and extract gel slice to a clean microcentrifuge tube
8. proceed with Promega Wizard Plus Gel and PCR kit, following manufacturers
protocol
9. elute sample in ~20 ul of elution buffer
STEP 8: Final preparation before Sequencing
1) qubit to determine concentration (2x 2ul)
2) OPTIONAL: multina/gel to determine size distribution of library
3) Use the qubit concentration as well as the size distribution to calculate the molarity of
the final library
i.
We have an excel calculator to help with this
4) Adjust the concentration so that you have one of:
a. 10 nM – best
b. 4 nM – good
c. 2 nM – absolute minimum
5) Ready for sequencing!
APPENDIX & RANDOM NOTES
General tips and tricks
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use multichannel pipettes where possible
o transfer master-mixes to 8-well strips so you can use the multichannels
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ampure XP cleanup
o accurate pipetting is critical when using ampure XP beads
o it is better in increase the total sample volume so that small changes in
pipetting accuracy have less of an effect
o use an aspirator pump for discarding supernatants during cleans. It is much
faster than pipetting out solutions, and leaves less behind
Sample preparation
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DNA samples should be ideally high molecular weight genomic DNA
o Run a subset of samples on a gel to check for consistency
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DNA should be free of RNA
o Use RNase somewhere in the DNA extraction
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DNA should be quantified by Qubit
o Nanodrop readings are not accurate for determining DNA concentration and
should not be used in any next generation sequencing protocol
o It is recommended to ensure high 260/280 ratios (>1.8 ideally), and this can
only be done on a spectrophotometer such as a nanodrop.
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DNA starting concentration
o The amount of DNA per sample must be standardized going into the DNA
digestion step
o The amount of DNA used will likely be influenced by the availability of DNA
that you have access to
o You should aim to have 100-500 ng per sample
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We have got the protocol working for much lower DNA
concentrations (10ng and <1ng per sample), however, it is likely to
more variable.
Ligation and barcodes
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it is important to check barcodes are compatible using the “barcode diversity
calculator” before starting this step
o only critical if running a small number of samples
o if using full 32 barcodes (see below), it is fine
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basic 32 barcode setup (see plate map below)
o note: this is not the entire set of barcodes used in the Pererson et al 2012
paper,
o these barcodes have been chosen specifically to allow you to move directly
from digestion to ligation without a cleanup in between
o using these barcodes with all indexes will allow multiplexing of up to 384
samples (the full set allows 576)
o if more than 32 samples, simply repeat distribution of some/all initial 32
barcodes as needed (you will need to ensure the indexes added in PCR step
differentiate samples that have the same barcode)
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***numbers represent barcodes order from Peterson el al 2012
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Double size selection using ampure beads
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double size selection refers to the use of beads to first bind high Mw DNA (>~800bp), after which the supernatant is collected, and a second bead bind is used to
collect >250-bp fragments
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should result in an enrichment of fragments in range of 250-800-bp
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this step is included in the protocol to help in a few ways
o It removes high and low Mw DNA that will not be sequenced regardless
o It enriches for DNA in approximately the correct size range prior to PCR, so
that when samples are PCRed, amplification of relevant fragments takes
place
o It helps in the normalization of samples that might differ in quality
PCR
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PCR is performed to extend the ligated RAD product, and add additional PCR-based
indexes onto the MspI end of the sequence
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P2 primers: It is important to ensure that there is diversity among P2 primer indexes
if running more than 32 samples
o if running 32 samples or less, use P2 primer 1 only
o if running > 32 samples, I would use at least the first 6 P2 primers.
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This ensures there is plenty of diversity among P2 indexes, which will
help in the demultiplexing step
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PCRs are performed using the KAPA KiFi Real Time amplification mix, which allows
detection of library amplification in real time on a qPCR machine
o This is not essential, however, it allows you to ensure that every sample gets
amplified, and allows toy to stop amplification as soon as a sufficient number
of cycles is reached
o This is better than titrating PCR cycles and running on gels as previously done
Pooling libraries
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the aim here is to pool all samples in eqimolar concentrations to ensure all samples
get sequenced equally/evenly
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we do this by combining the same amount (ie. ng) per sample into a single tube
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pooling approach will depend on how much DNA you have across all samples
o high concentrations – add a nominal amount, eg. 10-20 ng from each sample,
to a single tube.
o Low concentrations – determine what the lowest concentration sample is,
and multiply the volume by the concentration/ul to determine the total ng in
that sample. Then, add all of that sample to the pooling tube, and then add
the same amount of DNA from all other samples to the tube. You will
therefore use the entire sample with the lowest concentration.
Size selection of libraries
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libraries need to be size selected to reduce the number of unique reads sequenced
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the size of DNA fragments that you aim for ultimately depends on a number of
factors, including
o size of the genome
o number of samples you want to multiplex
o the number of sequencing reads you expect to have
o number of loci you are aiming for
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most of the time, these are unknown variables that will need to be worked out the
hard way, ie. by doing a trial sequencing run.
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As a general rule, as a genome size increases, so to does the number of unique RAD
fragments. Therefore, for bigger genomes, a smaller size range of fragments should
be selected.
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If no reference genome is available, search the literature to see if there is any
information about predicted genome size. A useful place to start is to see if there is a
c-value available
Figure: example of making “super-wells” by using electrical tape to join multiple wells
together. This allows the formation of a much larger well to add the pooled library to for
size selection
Figure: Example of gel showing excised libraries and a double-size selected and PCRed
library. There was two gel cuts made at approximately 400-600 and 600-800bp. However,
the size of gel cuts are species/genomic size dependent.