TITLE: PotatoMASH library construction
AUTHORS: Maria de la O Leyva Perez, Lea Vexler, Dan Milbourne

[DESCRIPTION]
We have developed PotatoMASH (Potato Multi-Allele Scanning Haplotags), a novel low cost, genome-scanning marker platform. We designed a panel of 339 multi-allelic regions placed at 1 Mb intervals throughout the euchromatic portion of the genome. These regions were assayed using a multiplex amplicon sequencing approach, which allows to genotype hundreds of plants at a cost of €5/sample.
This protocol describes the library construction part of the method. PotatoMASH libraries are made to be sequenced in Illumina sequencing platforms.


[STEPS]
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1. 
Quantification with PicoGreen
In the preparation for the library construction, the first step is to quantify the template DNA. We used Quant-iT™ PicoGreen ® dsDNA assay kit (Invitrogen, P7589)as detailed below: 

The Quant-iT™  PicoGreen kit should be stored in 4 °C in the dark. An hour before take the PicoGreen kit out from the fridge and place in room temp, protect the PicoGreen reagent from light by covering the tube with aluminium foil. The Lambda DNA standard should be always kept in 4°C. 

Notes: 
Step duration is estimated for one plate. However, the duration for each step and sub-steps of this protocol is not necessarily the result of multiplying the duration of all steps by the number of plates to process. For example, in this step 1 (Quantification with Picogreen) the duration should be multiplied by the number of plates to process only in steps 1.3, 1.4, 1.6, 1.8 and 1.10.
It is advised to read the manufacturer's protocol   


SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.1. Prepare the Lambda DNA standarts in a serial dilution: 
Note: Vortex and spin down between each dilution.
  
     Concentration of standard (λ)      Volume of 1X TE      Volume and concentration of Lambda      Final DNA Concentration  PicoGreen Assay        2000   ng/ml      1470   μL       30   μL of   100 μg/mL λ       1000   ng/ml        200   ng/ml      1350 μL      150 μL of   2000 ng/mL λ       100   ng/ml        20   ng/ml      1350 μL      150 μL of   200 ng/mL λ       10   ng/ml        2   ng/ml      1350 μL      150 μL of   20 ng/mL λ       1   ng/ml   
 
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.2. In 50 ml tube Prepare 1X TE as following TWICE (the second load is for the PicoGreen working solution - in step 1.5 ): 
 
     Reagent      96-well plate      4 x 96-well plates        20X TE      500 µl      2 ml         water      9.5 ml      38 ml   
 Mix by vortex.
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.3. Dilute DNAs in black 96-well plate for quantification: 
In columns 1-11: 
Pipette 98 µl of 1X TE in each well
Add 2 µl of each DNA sample (multichannel pipette recommended).
Mix by pipetting.
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.4. In column 12 : Fill in with 100 µl of the standards as follows: 
 
     12A      2000   µg/ml   λ        12B      200   µg/ml   λ        12C      20   µg/ml   λ        12D      2 µg/ml   λ        12E      1X   TE        12F      Empty        12G      Empty        12H      Empty   
 
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.5. Prepare the working solution:
 
     Reagent      96-well plate      4 x 96-well plates        1X TE (from step)      9.9 ml      39.6 ml        PicoGreen reagent      100 µl      400 µl   

Protect the working solution from light by covering the tube with aluminium foil.

SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.6. Add 100 µl of working solution in each well.
Use the same tips to mix by pipetting.
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.7. Incubate for 2 to 5 minutes at room temperature, protected from light.
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.8. Measure by using a fluorescence microplate reader and standard fluorescein wavelengths (excitation ~480 nm, emission ~520 nm). The instrument’s gain should be set to the well containing the highest DNA concentration (12A)
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.9. Calculate template DNA concentrations:
Subtract the fluorescence value of the reagent blank (12E) from that of each of the samples. Use corrected data to generate a standard curve of fluorescence versus DNA concentration. Determine the DNA concentration of the sample from the standard curve and multiply by the dilution (x100)
SECTION: Organize your samples in 96 well plates, quantitate template DNA, and normalize at 20ng/uL.
1.10. Template DNA Normalization:
Make dilutions in a new plate or add nuclease free water according to each sample concentration to bring all samples to 20 ng/µl.
SECTION: PotatoMASH PCR1 (multiplex PCR of 339 loci)
2. This step amplifies all PotatoMASH loci and adds Illumina sequencing primer tags R1 and R2.


     
PotatoMASH primers can be downloaded as supplemental data of the publication: Leyva-Pérez, M.O.; Vexler, L.; Byrne, S.; Clot, C.; Meade, F.; Griffin, D.; Ruttink, T.;  Kang, J. and Milbourne, D. PotatoMASH - a low cost, genome-scanning marker system for use in potato genomics and genetics applications. Please cite this work if using this protocol.

If you decide to skip the library normalization (step 5) use the thermocycling conditions in 2.4 B

Prepare the PCR cocktail the same day for the number of plates to be processed.

Additional note: step duration is estimated for one plate. However, the duration for each step and sub-steps of this protocol is not necessarily the result of multiplying the duration of all steps by the number of plates to process.
SECTION: PotatoMASH PCR1 (multiplex PCR of 339 loci)
2.1. Prepare the PCR1 Cocktail: keep on ice   
Note: It is possible to prepare enough cocktail for all the plates planned to do the same day
     Reagent      1 sample      Cocktail for 96-well plate         Qaigen Master Mix (2x)      3.5 µl      350 µl        PotatoMASH primers      1.4 µl      140 µl        nuclease free water      0.1 µl      10 µl        DNA (20ng/µl)      2 µl      -        Total      7 µl      500 µl   (5 µl   per sample)   
 Note: Advised order to prepare the cocktail: 1st pipette the water, 2nd add the primers, 3rd the Qiagen Master Mix.

Gently mix by pipetting.
SECTION: PotatoMASH PCR1 (multiplex PCR of 339 loci)
2.2. Dispense 5 µL of the PCR1 Cocktail into each well

Note: it is advised to use 100 µl electronic pipette with dispenser function to apply the 5 µl of the PCR1 Cocktail to the plate with a 100 µl filter tip. A single tip can be used for all the plate.
SECTION: PotatoMASH PCR1 (multiplex PCR of 339 loci)
2.3. Apply 2 µL  of the DNA (20ng/µl) with 10 µl filter tips, 
Mix by pipetting.
Seal the plate.
Spin down in a mini plate spinner for 10 sec.
SECTION: PotatoMASH PCR1 (multiplex PCR of 339 loci)
2.4. In a gradient thermocycler with heated lid, run PCR1 as follows:
A - Follow when performing library normalization (step 5)
 
                  Cycles  Temp  Time  Temp  Time  Temp  Time    1x  90 °C  15 min            8x  95 °C  30 sec  0.2°C /sec ramp down to 57°C  30 sec  72 °C  2 min    16x  95 °C  30 sec  65 °C  30 sec  72 °C  30 sec    1x  8 °C  ∞          
 
B- Follow when skipping library normalization (step 5):

                  Cycles  Temp  Time  Temp  Time  Temp  Time    1x  90 °C  15 min            8x  95 °C  30 sec  0.2°C /sec ramp down to 57°C  30 sec  72 °C  2 min    16x  95 °C  30 sec  65 °C  30 sec  72 °C  30 sec    1x  8 °C  ∞          

 Note: PCR1 product can be stored at 4 °C 1-2 days or longer at -20 °C before dilution.
SECTION: Dilute PCR1 product to be used as template for PCR2
3. Note that it is recommended to dilute PCR1 product the same day PCR2 will be performed.

Centrifuge the PCR1 product plate to make sure all volume is down (e.g. for 3 min at 3000 RPM 15 °C )
SECTION: Dilute PCR1 product to be used as template for PCR2
3.1. Add 100 µL  of nuclease free water to each well to dilute 1:15.
Mix by pipetting up and down 30 times.
Note: it is convenient to use a multichannel 100 µl electronic pipette with function 'pipette and mix' set up to dispense 100 µL and mix 40 µL volume 30x.

SECTION: Dilute PCR1 product to be used as template for PCR2
3.2. Spin down for 10 sec in a mini plate spinner.
Diluted plates can be kept at 4 °C meanwhile preparing the cocktail for PCR2
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4. PCR2 adds Illumina capture adapter P5 and P7 and barcode indexes that identify each sample by well and by plate.

      

Primers i5 provide with well-specific barcodes. There is a panel of 96 primers. We use the same panel for each plate. 
Primers i7 provide with plate-specific barcodes.

The 96 i5 primers and 10 i7 primers can be downloaded as supplemental data of the publication: Leyva-Pérez, M.O.; Vexler, L.; Byrne, S.; Clot, C.; Meade, F.; Griffin, D.; Ruttink, T.;  Kang, J. and Milbourne, D. PotatoMASH - a low cost, genome-scanning marker system for use in potato genomics and genetics applications. Please cite this work if using this protocol.

Here we provide with additional i7 primers to multiplex more plates (plate 11 to plate 19)
  
By using 96 i5 barcodes in combination with 19 i7 barcodes, they can be indexed up to 1,824 samples to be joined in a single library.
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4.1. Prepare the PCR2 Cocktail: keep on ice. on ice 

 
     Reagent      1 sample      Cocktail for 96-well plate        Quiagen Master Mix (2x)      5 µl      500 µl        i5 10 µM primers      1 µl      -        i7 10 µM primers      1 µl      100 µl        DNA (20ng/ul)      3 µl      -        Total      10 µl      600 µl   (6 µl   per sample)   
Notes: Since i7 primers are plate-specific, you will need to prepare a different PCR cocktail for each plate. Advised order to add reagents: 1st apply the i7 primers, 2nd the Qiagen Master Mix. Mix by pipetting and spin down 10 sec.
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4.2. Dispense 6 µL of the PCR2 Cocktail into each well

Note: it is advised to use 100 µl electronic pipette with  the '(Disp.)' function to dispense 6 µl of the PCR2 Cocktail to each well in the plate with one 100 µl filter tip. A single tip can be used for all the plate.
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4.3. Add  1 µL of the well-specific i5 primers.

Note: it is advised to use the manual (not electronic)10 µl multi-channel pipettor to add the 1 µl of the well- specific i5 primers from the plate were the 96 primers are to your PCr2 plate. That way is easier to place each of the 96 i5 primers in each well position and to reduce the chance of mistakes. Make sure the 1 µL is delivered by pipetting up-down a couple of times.
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4.4. Add 3 µL  of the diluted PCR 1 product and mix by pipetting.
Seal the plate.
Spin down in a mini plate spinner for 10 sec.
SECTION: PotatoMASH PCR2 (Adds dual barcodes)
4.5. Run PCR as follows:
 
                    Step  Cycles  Temp  Time  Temp  Time  Temp  Time    Initial   Denaturation    90 °C  15 min            Denaturation   & Annealing  10x  98 °C  10 sec  65 °C  30 sec  72 °C  30 sec    Elongation  1x  75 °C  5 min            Store  1x  10 °C  ∞          
 Note: The libraries (PCR2 product) can be stored at 4 °C 1-2 days or longer at -20 °C.
SECTION: Library normalization (optional)
5. This step is optional. By having normalized the template genomic DNA prior to PCR, we minimized the differences of efficiency in each PCR. Homogeneous template DNA quality also minimizes differences in PCR yield in each well. However, if you are genotyping sets of DNA extracts within the same plate of very variable quality, we recommend to follow the complete protocol including library normalization step.
When performing this step be aware to have followed step 2.4A and proceed at later stage with step 6.2A.
When skipping this step be aware to have followed step 2.4B and proceed at later stage with step 6.2B.

In this step we normalize the PCR2 product in each well down to ∼25 ng in 20 µL (∼1.25ng/µL).

  

As an additional benefit, it also provides with a first purification step: the binding buffer provides low pH condition causing the positive surface charge of coating of the library normalization plates to bind the negatively charged nucleic acid backbone. Proteins and other contaminants (such as short oligonucleotide primers) are not bound and are simply washed away.

Tip: In this step, it is possible to use unfiltered tips, since the libraries are already prepared and barcoded, therefore, the risk of contamination is lower.
It is advised to read the manufacturer's protocol (Applied Biosystems, A105100)   

SECTION: Library normalization (optional)
5.1. Before starting:
Centrifuge the PCR2 product plate to make sure all volume is down (e.g. 3 min at 3000 RPM 15 °C )

SECTION: Library normalization (optional)
5.2. Binding:
With a multichannel pipette, add 10 µL binding solution (the same volume of the PCR product) into the PCR2 plate and mix completely by pipetting up and down. With the same tips, transfer 10 µl, twice, to the normalization plate. 

Advice: Release on the upper part of the well, careful to not interrupt the coating by scratching the well sides.
 
Seal the plate and spin down in a mini plate spinner for 10 sec. 
Incubate in 15 Room temperature for 1 hour
Transfer back the amplicon/Binding Buffer mixture to the original PCR 2 plate.

After binding is complete, the mixture can be stored at –20ºC for up to 30 days to recover more libraries later if needed. Again, be careful no to scrape the well sides with the tips during aspiration. To use 10 µL (transfer 10 µL twice), helps to avoid scrapings.
SECTION: Library normalization (optional)
5.3. Washing:
Add 50 µL of wash buffer. Mix by pipetting up and down 35 µl 3 times. 

Note: it is convenient to use a multichannel 100 µl electronic pipette with function 'pipette and mix' set up to dispense 50 µL and mix 35 µL volume 3x. Then you can press the pipette button again to use the same tips to aspirate and discard the washing buffer.

Use the same tips to aspirate and discard the wash buffer.
Invert and tap the plate on a paper towel to get rid of the remaining droplets of liquid. 
SECTION: Library normalization (optional)
5.4. Elution:
Elute the libraries with 20 µL of elution buffer (pH 8.5–9.0). Mix by pipetting up and down 10 µl 5 times.
Incubate at room temperature 5 min.
The plates with the eluted DNA can be stored at 4°C (short-term storage) or –20°C (long-term storage). 
SECTION: Library normalization (optional)
5.5. Store in 4 °C until library pooling.

SECTION: Pooling libraries, Concentration - Purification
6. Pooling sub-libraries:
At this stage, the libraries are barcoded and even normalized. We can pool a full plate (96 samples) into one 2mL tube.

  

We also refer to each of the pools as 'pooled sub-library'. 
It is convenient to keep the plate on a cooling holder or on ice .
SECTION: Pooling libraries, Concentration - Purification
6.1. Before starting:
Spin down the plates.
Label one tube per plate on ice .

SECTION: Pooling libraries, Concentration - Purification
6.2. A - Follow if you have performed library normalization (step 5):
Transfer 15 µL of each sample per 96-well plate into one 2mL tube.

B- Follow if you have skipped the library normalization (step 5):
Transfer 5 µL of each sample per 96-well plate into one 1.5mL or 2mL tube.

Note: we advise to use a 200uL electronic pipet with multi-aspirate function to take 15 µl x 12 times (pool by rows) and deliver in the same tube. All can be done with 1 filter tip. Tubes can be kept at 4 °C or on ice  for short term prior to Purification steps the same day or at -20°C for longer term.

SECTION: Pooling libraries, Concentration - Purification
7. Concentration - Purification:
In this step we use the purification kit to purify, concentrate and reduce the volume of each pooled sub-library from 0.25 or 1.5 mL to 42 µl by using the Qiaquick PCR Purification kit (Qiagen, 28104) and extra binding buffer (PB buffer, Qiagen, 19066), following manufacturer's guidelines with slight adjustments:

Note: It is advised to read the manufacturer's protocol    or   

SECTION: Pooling libraries, Concentration - Purification
7.1. Before starting:
Spin down the tubes with the pooled sub-libraries. 
Label 15 ml tube (in case of normalized libraries, protocol A) or one 2mL tube (in case of normalized libraries, protocol B) for each pooled sub-library for the binding step (7.2)
Prepare collection tubes with a QIAquick Spin column inserted for each sub-library for the binding and washing step (7.2-7.3)
Prepare same number of 1.5 ml tubes for the elution step (7.4)
SECTION: Pooling libraries, Concentration - Purification
7.2. Binding:
A- Follow if you have performed library normalization (step 5):
Prepare a 15 ml tube for each pool with 7.5 mL of PB
The ratio is 5 volumes of Buffer PB to 1 volume of the PCR product.
Add the 1440 µL of the pooled sub-libraries
B- Follow if you have skipped the library normalization (step 5):
Prepare a 2 ml tube for each pool with 1.2 mL of PB
Important: Make sure that the non-normalized pool is well mixed by vortexing a few seconds if you didn't so right after pooling the libraries. It is important because we will use only the half of the volume. The remaining volume can be stored at -20 °C in case is needed.
Add the 240 µL of the pooled sub-libraries

From now on, all steps are in common for protocols A and B:

Vortex for 5 sec after mixing each pool with the PB and again after having processed all of the tubes. 
Pipette 600 µL of the PB-PCR product mix into the QIAquick Spin column.
Centrifuge for 30 sec at 13,000 RPM and discard the flow-through.
Repeat the last 2 steps until all the PB-PCR product mix in the tube is finished. 
SECTION: Pooling libraries, Concentration - Purification
7.3. Washing:
Add 750 µl Buffer PE to the QIAquick Spin column,
Centrifuge for 60 sec at 13,000 RPM and discard the flow-through.
Centrifuge for 60 sec at 13,000 RPM to dry.
SECTION: Pooling libraries, Concentration - Purification
7.4. Elution:
Transfer the  QIAquick Spin column to a new 1.5 ml tube.
Elute with 44 µL Elution Buffer
Incubate in 4 Room temperature for 1-5 min. 
Centrifuge for 1 min at 13,000 RPM. 
SECTION: Pooling libraries, Concentration - Purification
7.5. Quality check (optional):
Use 2 µL of the eluted pooled libraries to check quality with nanodrop. 
Expected concentration values: 30 – 80 ng/µl. 
Expected ratio A260nm/A280nm: 1.8 – 2,1
Expected ratio A260nm/A230nm: 2-2.5 
SECTION: Size selection
8. This step is meant to remove PCR1 primer-dimer amplified products in PCR2 and other unwanted secondary products from the PotatoMASH libraries. All PotatoMASH core primers are designed to produce libraries of around 300bp. Therefore, electrophoresis in agarose gel is a low-cost and easy way to separate them.

Note that we usually perform this step the same day we perform step 7, but probably the concentrated libraries can be stored at 4°C for a day or at -20°C for longer term.
SECTION: Size selection
8.1. Prepare 1.2 % standard low melting point agarose gel in 0.5x TBE (TAE buffer can be also used).
For DNA visualization, we use GelRed dye applied to the agarose gel before polymerizing.
Make sure the wells of the gel are wide-deep enough to hold 48 µL of sample. That can be achieved, for example, by covering three 7 mm. wide x 1mm thick comb teeth (3 wells) with scotch tape to make a wider well of 2.5 cm x 1mm thick. We recommend to leave one normal size well empty between different pools.

Tip: Around 200mL of agarose are used for a gel 14cm long and 23 cm wide 7mm thick to get the wells deep enough to hold 48 µL. Try to get the gel only  thick enough to hold the sample but not more. The more thick is the gel, the more agarose you will have to clean up in the next step.
SECTION: Size selection
8.2. Add 8 µL of loading buffer 6X to the 40 µl of each of the tubes with the concentrated libraries. Mix by pipetting. 
SECTION: Size selection
8.3. Load the sub-libraries onto the big wells. 
Load 6 µL of p100 DNA ladder (e.g. NE Biolabs N3231) into at least one of middle wells in between. We do this to confirm that our main product is of the expected size and help us to spot it. 
SECTION: Size selection
8.4. Run in 50 V for 1 hour and then run for 90 min with 100 V. 

Note: Settings used for a gel 14 cm long and 23 cm wide 0.7 cm thick, 200-230mL of agarose.
SECTION: Size selection
8.5. Use a transilluminator UV (recommended to use low intensity like 70%) to visualise and excise the band around 300bp. Use a clean scalpel and place it in a 2 mL tube.
 
   

 

SECTION: Size selection
8.6. Weight the gel slices and record the weights (the weight will be important for next step, gel purification)
At that point, the gel slice can be stored in a closed tube at  4 °C for a couple of days if necessary. 

SECTION: Gel purification
9. Purify the slice using the Wizard® SV Gel and PCR Clean-Up System Kit (Promega A9281) following manufacturer's guidelines:

Note: It is advised to read the manufacturer's protocol   
SECTION: Gel purification
9.1. Before starting:

Heat a thermoblock  to 65 °C
Prepare collection tubes with SV mini-column for each sub-library for the extraction step (9.3)
Prepare same amount of 1.5 mL tubes for the elution step (9.5)
SECTION: Gel purification
9.2. Dissolving the gel
Add 10 µL of membrane binding solution per 10mg of gel slice.
Warm up to 50-65 °C vortexing from time to time until the agarose completely dissolves.
Vortex or mix by inversion. 
SECTION: Gel purification
9.3. Extraction of PCR product (libraries) from the dissolved gel:
Transfer 650 µL of the dissolved gel mixture to the SV mini-column and a collection tube
Incubate for 1 min in 65 Room temperature. 
Centrifuge for 1 min at 14,000 RPM (16,000 x g) and discard flow-through.
Repeat until all the dissolved gel mixture have passed through the SV mini-column. 
SECTION: Gel purification
9.4. Washing:
Wash with 700 µL of washing solution
Centrifuge 1 min at 14,000 RPM (16,000 x g). Discard flow-through
repeat with 500 µl of washing solution. Discard flow-through
Centrifuge 5 min at14,000 RPM (16,000 x g) to dry the column.
SECTION: Gel purification
9.5. Elution: 
Transfer the mini-column to a 1.5 ml tube.
Elute with 50 µL Nuclease-free water (pre warm 65 °C)
Incubate for 1-5 min in 4 Room temperature.
Centrifuge 1 min at 14,000 RPM
SECTION: Gel purification
9.6. Quality check:
Use 2 µL of the eluted pooled libraries to check quality with nanodrop. 
Expected concentration values: 5 – 25 ng/µl. (The higher yields are expected for non-normalized libraries. Note that nanodrop measurements are not accurate for very small concentrations)
Expected ratio A260nm/A280nm: 1.8 – 2,5
Expected ratio A260nm/A230nm: 0 – 1

Store in 4 °C until next step.


SECTION: Purification with Magnetic AMPure XP beads
10. The last purification step is performed with Magnetic AMPure XP beads (Beckman C. A63881) on each of the pooled sub-libraries and by using a magnetic stand for 1.5mL microcentrifuge tubes.
This step is necessary to achieve the level of library purity needed for posterior Illumina sequencing. Note that the Expected ratio A260nm/A230nm of the pooled sub-libraries after gel purification are not good.

As an additional benefit, the  method  utilizes  a  double-size  selection  steps that will further purify remaining traces of PCR products larger or smaller than 300bp that weren't properly separated during the size selection by agarose gel.
In the first selection, beads will bind in higher proportion to fragments larger than 400 bp. Then we recover the clear supernatant with fragments smaller than 400 bp. In the second selection, beads will bind in higher proportion to  fragments larger than 200 bp and by discarding the clear supernatant we will discard fragments smaller than 200 bp. 

 Note: It is advised to read the manufacturer's protocol    

SECTION: Purification with Magnetic AMPure XP beads
10.1. Before starting:
Prepare  5 mL 85 % Ethanol.
Magnetic Ampure XP beads stock to be stored 4 °C 
Resuspend the Ampure XP beads stock Suspension (usually in a bottle) carefully until complete homogenization.

Tip: It is convenient to separate from the bottle the volume of ampure beads you'd need for all tubes into a small tube, which is easier to vortex and get homogeneus every time is needed.

Spin down the sub-libraries (if they were in the fridge or freezer) and measure the volume of library sample (∼ 46 µl). 

SECTION: Purification with Magnetic AMPure XP beads
10.2. First purification:
Vortex again the AMPure beads.
Add 0.5 x volumes of AMPure beads (∼ 23 µl) of the sample volume (∼ 46 µL) and mix by pipetting 10 times.
Vortex for 30 sec.
Incubate in room temperature for 1 min. 
Place the reaction tube on a magnetic stand for 1-5 min to allow the AMPure beads to settle to the magnet until the supernatant is clear. 
Recover the clear supernatant, measure and record the volume. Place in a new tube. 

Note: it is advised to use an electronic pipet with function 'manual (man.)' to pipette very slowly and measure the recovered volume.
SECTION: Purification with Magnetic AMPure XP beads
10.3. Second selection:
Vortex the AMPure beads again.
Add 0.6x volumes of AMPure beads (∼ 42 µL) to the recovered supernatant (∼  70 µL). Mix by pipetting 30 times. 
Vortex again for 1 min. incubate at room temperature for 1 min.
Place the reaction tube on the magnetic stand for and allow the AMPure beads to settle to the magnet. 
Remove and discard the clear supernatant 
Notes: It is ok to leave a droplet at the bottom, it will wash in the next step with the ethanol. Careful not to aspirate any AMPure beads.
SECTION: Purification with Magnetic AMPure XP beads
10.4. Wash with 85 % ethanol 
While the reaction tube still on the magnet, add 180 µL of 85% ethanol 
incubate at 4 Room temperature for 30-60 seconds. 
Carefully Remove and discard the ethanol supernatant by pipetting. 
Leave the tube open for 30 sec to dry.
SECTION: Purification with Magnetic AMPure XP beads
10.5. Elution: 
Remove the reaction tube from the magnet
 Add 30 µL of Nuclease-free water. 
Mix the total elution volume by pipetting 30 times to resuspend the beads
Incubate at 4 Room temperature for 1 minute.
Place the reaction tube on the magnetic stand for 1 min or more and allow the Ampure beads to settle to the magnet.
Carefully transfer by pipetting the eluted pooled sub-library (∼ 27 µl) into new tubes.

SECTION: Purification with Magnetic AMPure XP beads
10.6. Quality check:
Use 2 µL of the eluted pooled sub-libraries to check quality with nanodrop. 
Expected concentration values: 7 – 35 ng/µl. (The higher yields are expected for non-normalized libraries. Note that nanodrop measurements are not accurate for very small concentrations)
Expected A260nm/A280nm ratio: 1.8 – 2.2
Expected A260nm/A230nm ratio: 1.5 – 2
SECTION: Sub-libraries quantitation and final pooling into a single library
11. Quantitation:
Sub-libraries quantitation using Qubit™ dsDNA BR Assay Kit (Thermofisher Q32853) following manufacturer's guidelines   
Given the expected yield, we recommend to use 3-5 µL of each sub-library to quantify them with the Broad spectrum assay kit. Alternatively, The HS (high sensitivity) assay kit can be used.

Pooling sub-libraries into a single library:
Calculate the volume from each sub-library to be pooled so there is the same amount in ng of each sub-library in the volume of the final library. Pool the sub-libraries and mix by pipetting.

  

Make sure that your final PotatoMASH library meets the sample requirements for Illumina sequencing of pre-made libraries (PCR free library), which usually are >0.5-1ng/ µL (quantified by Qubit), >20 µL or more volume depending of the amount of sequencing data required.
Sequencing service companies usually offer demultiplexing of libraries and for that the ask the barcode of each sublibrary. Here is an example of how to code the sublibraries:

  
You can also demultiplex the data yourself and you will need the i5 and i7 barcodes of each sample. The tags are written in the name of each primer, but here you have them in a table:

  

This method has successfully genotyped 765 samples in a whole lane of  Illumina HiseqX platform using 50% PhiX, obtaining paired-end 150nt reads (56Gb of data). The libraries have been also proven to be successful for sequencing in NovaSeq 6000 platform.

  
SECTION: Library quality check (optional)
12. Library quality check (optional) by 'Illumina PCR' (PCR with known primers which anneal to Illumina capture adapters P5 and P7).
This is an alternative method to check the quality of your libraries without the need to use bioanalyzer or fragment analyzer.
It can be performed only on the Final library or to the sub-libraries before pooling them, in case you would prefer to further purify one of them before pooling.
The primers will anneal to any PCR product containing P5 and P7, so you can detect if you still have too many small fragments in your sub-libraries or final library.
Before starting:
Order the primers:
P5 primer: 5'-AAT GAT ACG GCG ACC ACC GA-3'
P7 primer: 5'-CAA GCA GAA GAC GGC ATA CGA-3'
SECTION: Library quality check (optional)
12.1. Dilute the library/libraries to test to 0.1 ng/µL. You will only need 1µL of this dilution.
SECTION: Library quality check (optional)
12.2. Prepare the Illumina PCR cocktail:
 
     Reagent      1 sample      7 samples        Qiagen Master Mix (2x)      5 µl      35 µl        Illumina primers 10 µM (mix of both)      1 µl      7 µl        H2O      3 µl      21 µl        library (0.1 ng/ul)      1 µl      -        Total      10 µl      63   µl (9 µl per sample)   
 Distribute 9 μL of the cocktail per 0.2mL PCR tube and add 1μL of your library at 0.1 ng/μL. Mix by pipetting and /or spin down the tubes.
SECTION: Library quality check (optional)
12.3. Run PCR as follows:
 
                Step  Cycles  Temp  Time  Temp  Time    Denaturation    90 °C  1 min        Annealing & Elongation  16x  95 °C  15 sec  63 °C  45 sec    Store  1x  10 °C  ∞      
Be aware that the optimal elongation temperature for the polymerase in the Qiagen multiplex master mix is 72°C and not 63°C. However, it works nicely anyway, and the annealing and elongation are done at the same temperature making the program shorter. Proper optimization is always welcome though.
SECTION: Library quality check (optional)
12.4. Run the PCR product (10 µL) mixed with 2 µL of loading buffer (6x) on 1.2 % agarose gel. Use ladder p100.

      
3 examples of libraries tested. Notice that brightness/contrast have been increased to highlight the unspecific bands
√: the library passed quality test made in fragment analyzer by the sequencing service. This library was made skipping the library normalization (protocol B).
*The libraries were set on hold by the service because they contained small fragments. However, the customer can ask these on hold libraries to be sequenced anyway and they were successful.
We don't have examples of libraries failing quality control to be rejected for sequencing. However, we advise to keep in mind that the risk of having too much unspecific libraries is to loose part of the sequencing data in non-informative data.