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ddRADseq Library Preparation Cheat SheetWritten by Steve Doyle ([email protected]) - Feb 2014

- simplified version of ddRADseq from Peterson et al 2012 PLoS One

- 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

- keep track of regents used, either when completing complete protocol or optimizing

particular steps

Sample Setup for up to 96 wells

- 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

- 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.

mailto:[email protected]

STEP 1: DNA digestion - see appendix notes on sample preparation

Preparation

- thaw on ice

o cutsmart buffer

- 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

- thaw on ice

o T4 DNA ligase buffer (note: this is sensitive to freeze/thaw cycles – it is best

that this is aliquoted)

- put T4 DNA ligase on ice

- 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

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



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 ***



T4 DNA ligase buffer 4

T4 DNA ligase 1

Adapter P2 (2 uM stock) 2

H2O 1

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

- 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

- make Ampure mix 1

- make Ampure mix 2

- 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)



Ampure XP 10

PEG buffer 40



Ampure XP 10

PEG buffer 10

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

- see appendix for notes on PCR

- 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

1. setup PCR master-mixes, making sure they differ in P2 primer if needed



KAPA HiFi Real Time master

mix

10

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

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)



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 librariesPreparation

- make Ampure mix

-

-

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


seconds









11. Once ethanol has evaporated, add 20 ul of H20 and pipette up and down 10X to mix,

making sure beads have resuspended


(# samples + 4 extra)

Ampure XP 10

PEG buffer 60

12. allow to incubate for at least 10 mins


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)

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

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


seconds









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


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

- 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 Sequencing1) 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

- use multichannel pipettes where possible

o transfer master-mixes to 8-well strips so you can use the multichannels

- 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

- DNA samples should be ideally high molecular weight genomic DNA

o Run a subset of samples on a gel to check for consistency

- DNA should be free of RNA

o Use RNase somewhere in the DNA extraction

- 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.

- 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

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

- 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

- 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)

***numbers represent barcodes order from Peterson el al 2012

Double size selection using ampure beads

- double size selection refers to the use of beads to first bind high Mw DNA (>~800-

bp), after which the supernatant is collected, and a second bead bind is used to

collect >250-bp fragments

- should result in an enrichment of fragments in range of 250-800-bp

- 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

- PCR is performed to extend the ligated RAD product, and add additional PCR-based

indexes onto the MspI end of the sequence

- 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.

This ensures there is plenty of diversity among P2 indexes, which will

help in the demultiplexing step

- 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

- the aim here is to pool all samples in eqimolar concentrations to ensure all samples

get sequenced equally/evenly

- we do this by combining the same amount (ie. ng) per sample into a single tube

- 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

- libraries need to be size selected to reduce the number of unique reads sequenced

- 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

- 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.

- 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.

- 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.

Documents

Web viewdetermine a suitable amount of DNA per sample to pool, keeping in mind to maximize the amount of DNA per sample as possible. ... load libraries into super-wells. You might