DSBEST v2.0 (Caroe et al. 2018)

Alicia Grealy

Published: 2023-09-25 DOI: 10.17504/protocols.io.j8nlkemwxl5r/v1

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Abstract

This bench protocol is based on the work of Caroe et al. (2018), for preparing shotgun libraries from double-stranded DNA, typically for ancient and degraded DNA. Please cite Caroe et al. (2018) if you use this bench protocol!

Before start

I have begun including a double-stranded positive control oligo (dsCL104) in the library preparation; this is a double-stranded form of the single-stranded oligo described in Gansauge and Meyer (2013). Always include no-template controls and any extraction controls.

Store PEG and ATP at -20 deg C and avoid repeated rounds of freeze thawing.

Store MyOne C1 Streptavidin beads at 4 deg C in a fridge.

Attachments

Caroe_et_al-2018-Methods_in_Ecology_and_Evolution.pdf Caroe_et_al_2018_Supp_Info.pdf DS_BEST_SCHEMATIC.pdf DS_BEST_Sequencing_expectations.pdf

Steps

Preparation

Note

Perform all reaction set-up steps in a reagent-only pre-PCR space inside a dedicated ultraclean environment. Add DNA and subsequent master-mixes to the reaction, and perform wash steps, in a separate pre-PCR space.

"Suit up" in this order: hair net, nitrile gloves, facemask, coveralls, gumboots, booties, second pair of gloves.

Prepare the space by decontaminating surfaces with 10% household bleach followed by 70% ethanol. UV irradiate pipettes and racks. Racks should be bleached between subsequent uses and UV irradiated.

Ensure ice is available. Thaw reagents on ice as needed. Keep enzymes on ice at all times. Do not vortex enzymes to mix but mix by flicking the tube gently. Pulse centrifuge all reagents before opening.

Label tubes.

A	B	C
Tube	Qty	For ...
1.5 ml Safelock Tube	10	5X SYBR
0.5 ml Safelock Tube	8	25 mM dNTPs
1.5 ml Safelock Tube	1	0.1 uM CL104_duplex
1.5 ml Safelock Tube	1	0.1 uM CL105_duplex 1/500
1.5 ml Safelock Tube	1	10 uM IS7
1.5 ml Safelock Tube	1	10 uM IS8
1.5 ml Safelock Tube	1	10 uM CL107
1.5 ml Safelock Tube	1	10 uM CL108
1.5 ml Safelock Tube	10	CL105_106 STD dilution series 10^11 - 10^2
15 ml Falcon Tube	1	TE Buffer
15 ml Falcon Tube	1	TET buffer
1.5 ml Safelock Tube	1	Oligo hybridisation buffer
1.5 ml Safelock Tube	1	Reaction enhancer
15 ml Falcon Tube	1	EBT buffer
0.2 ml Lo-bind PCR tube	2	Adapters (P5_DS1, P7_DS2)
0.5 ml Lo-bind PCR tube	1	Adapters mix (DS_adapter_mix)
0.5 ml Lo-bind PCR tube	1	Adapters mix dilution
0.2 ml Lo-bind PCR Tube	# of samples + 2	Reaction tubes
1.5 ml Lo-bind Tube	3	Step 19, Step 24, Step 27 master mixes
1.5 ml Lo-bind Tube	# of samples + 2	Combining library with PB buffer
1.5 ml Lo-bind Tube	# of samples + 2	Elution of library from spin column
QIAGEN MinElute PCR Purification Spin Columns	# of samples + 2	Purification of library
0.5 ml Lo-bind Tube	# of samples + 2	Final library
0.5 ml Lo-bind Tube	# of samples + 2	1/20 dilution of library
1.5 ml Safelock Tube	2	Assay A and B master mixes
8-strip optical qPCR Tubes	(((# of samples + 2)*2)+26)/8	Assay A and B

Prepare all necessary buffers and UV decontaminate where appropriate.

Note

Aliquot 5X SYBR into 500-ul batches and store at -20 deg C in foil. Aliquot dNTPs into 50-ul batches and store at -20 deg C.

A	B	C	D
Buffer	Reagent	Volume to add	Final concentration in solution
Oligo hybridisation buffer 10X	5 M NaCl	100 ul	500 mM
	1 M Tris-HCl	10 ul	10 mM
	0.5 M EDTA	2 ul	1 mM
	Ultrapure water	888 ul	na
Reaction enhancer	50% PEG-4000	500 ul	25%
	10 mg/ml BSA	200 ul	2 mg/ml
	5 M NaCl	80 ul	400 mM
	Ultrapure water	220 ul	na
TE Buffer	Ultrapure water	9.88 ml	na
(Exp. 1 year)	1 M Tris-HCl	100 ul	0.01 M
	0.5 M EDTA	20 ul	0.001 M
EBT	1 M Tris-HCl	100 ul	0.01 M
	100% Tween 20	5 ul	0.05%
	Ultrapure water	9.895 ml	na
TET buffer	1 M Tris-HCl	500 ul	0.01 M
(Exp. 1 year)	0.5 M EDTA	100 ul	0.001 M
	100% Tween 20	25 ul	0.05%
	Ultrapure water	49.375 ml	na
5X SYBR	10,000X SYBR	2.5 ul	5X
	DMSO	997.5 ul	na
	DMSO	4 ml	na
25 mM dNTPs	100 mM dATP	100 ul	25 mM
	100 mM dTTP	100 ul	25 mM
	100 mM dCTP	100 ul	25 mM
	100 mM dGTP	100 ul	25 mM

Before resuspending oligos, pulse centrifuge to collect the pellet at the bottom of the tube. Add the appropriate buffer (see Materials) and vortex thoroughly. Store at -20 deg C. Dilute out the working concentrations (below) and store at -20 deg C when not in use. Thaw on ice. Vortex and pulse centrifuge after each thaw. Before beginning library preparation, make sure you have enough of each working stock prepared!

Note

Note: Do not store oligos and adapters in the same box as enzymes or reagents!The standards should be diluted in a totally different space, such as a teaching lab to ensure it does not contaminate the lab at extremetly high concentration. Also take extreme care with the positive control oligo as it will become a template for library preparation!

A	B	C
Working stock	Reagent	Volume to add
10 uM CL104_duplex	100 uM CL104	50 ul
	TET buffer	450 ul
0.1 uM CL104_duplex	10 uM CL104	5 ul
	TET buffer	495 ul
0.1 uM CL104_duplex 1/500	0.1 uM CL104	1 ul
(i.e., 0.0002 uM)	TET buffer	499 ul
10 uM IS7	100 uM IS7	50 ul
	Ultrapure water	450 ul
10 uM IS8	100 uM IS8	50 ul
	Ultrapure water	450 ul
10 uM CL107	100 uM CL107	50 ul
	Ultrapure water	450 ul
10 uM CL108	100 uM CL108	50 ul
	Ultrapure water	450 ul
10 uM CL105_106_STD	100 uM CL105_106_STD	50 ul
	TET buffer	450 ul
10^11 copies CL105_106_STD	10 uM CL105_106_STD	10 ul
	TET buffer	592.25 ul
10^10 copies CL105_106_STD	10^11 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^9 copies CL105_106_STD	10^10 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^8 copies CL105_106_STD	10^9 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^7copies CL105_106_STD	10^8 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^6 copies CL105_106_STD	10^7copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^5 copies CL105_106_STD	10^6 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^4 copies CL105_106_STD	10^5 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^3 copies CL105_106_STD	10^4 copies CL105_106_STD	50 ul
	TET buffer	450 ul
10^2 copies CL105_106_STD	10^3 copies CL105_106_STD	50 ul
	TET buffer	450 ul

Pre-program the thermal cycler.

Prepare adapters

In a 0.2 ml Lo-bind PCR tube ("P5_DS1"), combine:

40 ul of 500 uM IS1 

40 ul of 500 uM IS3

10 ul Ultrapure water

10 ul 10X Oligo Hybridisation Buffer

Vortex and pulse centrifuge.

In a new 0.2 ml Lo-bind PCR tube ("P7_DS2"), combine:

40 ul of 500 uM IS2

40 ul of 500 uM IS3

10 ul Ultrapure water

10 ul 10X Oligo Hybridisation Buffer

Vortex and pulse centrifuge.

10.

Incubate both "P5_DS1" and "P7_DS2" in a thermal cycler:

95 deg C for 10 sec

Ramp down to 12 deg C at a rate of 0.1 deg C/sec

11.

Combine both "P5_DS1" and "P7_DS2" together in a 0.5 ml Lo-bind Safelock tube to give 200 ul of 100 uM "DS_adapter_mix".

Determine the DNA input amount and concentration of adapters required

12.

Measure the concentration of your DNA extract on the Qubit following the manufacturer's instructions.

http://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0017209_Qubit_4_Fluorometer_UG.pdf

If possible, run the DNA on a fragment analyser or gel electrophoresis to determine the fragment length distribution.

http://www.perkinelmer.com/Content/LST_Software_Downloads/LabChip_GX_User_Manual.pdf

2% Agarose Gel Electrophoresis

Note

For ancient DNA it is seldom possible to quantitate the DNA or examine fragment lengths. For these, assume the average fragment length is 40 bp.

13.

Calculate the molarity of the dsDNA ends . Use the calculator at:

http://nebiocalculator.neb.com/#!/dsdnaends

Citation

e.g., 1 ul of 0.1 uM CL104_duplexMass dsDNA input (ng) = 3.71 ngLength (bp) = 60 bppmol dsDNA ends = 2*(((ng-input /1000/1000)/((length 617.96)+36.04))100010001000) = 0.2 pmolInput ds copy number = ((pmol dsDNA ends /2)/1000/1000/1000/1000)*(6.022E+23) = 6.02E+10

14.

Calculate the pmol of adapters needed. This is approximately 10X the molarity of the dsDNA ends. Round up to the nearest 10 pmol.

This number is equal to the concentration of adapters needed in uM.

Dilute the adapters to the desired working concentration in EBT buffer.

Citation

e.g., for the CL104_duplex above, 0.2 x 10 pmol adapter is needed, i.e., 2 pmol. Rounding up to the nearest 10 pmol gives 10 pmol. The concentration of adapters needed 10 uM.

Note

The final concentration of the adapters in the ligation reaction (below) should be 0.2 uM. Usually do not put in fewer than 10 pmol adapter, even for very low template samples.

End Repair

15.

Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.

A	B	C	D	E	F
Reagent	V2 (reaction volume)	C1 (stock concentration)	C2 (concentration in reaction)	V1 (volume to add)	x _______ rxn
T4 DNA ligase buffer w/ 10 mM ATP (NEB)	40 ul	10 X	1 X	4 ul
T4 DNA Polymerase	40 ul	3 U/ul	0.03 U/ul	0.4 ul
T4 PNK	40 ul	10 U/ul	0.25 U/ul	1 ul
Reaction enhancer	40 ul	na	na	2.2 ul
dNTPs	40 ul	25 mM	0.25 mM	0.4 ul

16.

Aliquot 8 ul into a 0.2 ml Lo-bind PCR tube for each reaction.

17.

To make the total reaction volume up to 40 ul, add:

Up to 32 ul DNA to each sample reaction. Make up the remainder with Ultrapure water. Typically input 3x10^8 - 3x10^11 double-stranded molecules; 13 pg-14 ng of ca. 40 bp DNA--this is typically 20% of the extract.

1 ul of 0.1 uM CL104_duplex positive control oligo to the positive conrol reaction + 31 ul Ultrapure water. We are inputing 3.01x10^10 molecules of single-stranded CL104 into the library preparation.

32 ul of Ultrapure water to the no-template control reaction.

Vortex and pulse centrifuge.

18.

Incubate in a thermal cycler at:

   20 deg C for 30 min

   65 deg C for 30 min

   4 deg C hold

Adapter ligation

19.

Add 1 ul of the DS_adapter_mix DILUTION (e.g., 10 uM) to the finished end-repair reaction above. Vortex and pulse centrifuge.

20.

Combine the following in a 0.5 ml Lo-bind tube. Vortex and pulse centrifuge.

A	B	C	D	E	F
Reagent	V2 (reaction volume)	C1 (stock concentration)	C2 (concentration in reaction)	V1 (volume to add)	x _______ rxn
T4 DNA ligase buffer w/ 10 mM ATP (NEB)	50 ul	10 X	0.2 X	1 ul
PEG-4000	50 ul	50%	6%	6 ul
T4 DNA ligase	50 ul	400 U/ul	8 U/ul	1 ul
Ultrapure water	50 ul	na	na	1 ul

21.

Add 9 ul to each reaction. Vortex and pulse centrifuge.

22.

Incubate in a thermal cycler at:

20 deg C for 30 min

65 deg C for 10 min

4 deg C hold

Fill in

23.

Combine the following in a 0.5 ml Lo-bind tube. Vortex and pulse centrifuge.

A	B	C	D	E	F
Reagent	V2 (reaction volume)	C1 (stock concentration)	C2 (concentration in reaction)	V1 (volume to add)	x _______ rxn
Isothermal amplification buffer	60 ul	10 X	0.33 X	2 ul
dNTPs	60 ul	25 mM	0.33 mM	0.8 ul
Bst 2.0 Warmstart DNA polymerase	60 ul	8 U/ul	0.213 U/ul	1.6 ul
Ultrapure water	60 ul	na	na	5.6 ul

24.

Add 10 ul of the above mix to each reaction. Vortex and pulse centrifuge.

25.

Incubate in a thermal cycler at:

65 deg C for 15 min

80 deg C for 15 min

4 deg C hold

Purify library

26.

Purify the libraries using a QIAGEN MinElute PCR Purification kit.

Note

Be sure to store the silica columns at 4 deg C.

http://www.qiagen.com/au/resources/resourcedetail?id=e24a0a3a-e9cb-4180-a6b4-202c527b924c&lang=en

Briefly...

27.

Combine 300 ul of PB Buffer with each reaction in a 1.5 ml Lo-Bind tube. Vortex and pulse centrifuge.

28.

Transfer the mixture to a MinElute Silica Spin Column (purple) placed inside a collection tube. Centrifuge for 1 min at 13,000 rpm in a bench-top centrifuge. Discard the flow-through.

29.

Add 700 ul of PE Buffer to the column. Centrifuge for 1 min at 13,000 rpm in a bench-top centrifuge. Discard the flow-through.

30.

Repeat Step 29.

31.

Centrifuge one more time (dry) for 1 min at 13,000 rpm in a bench-top centrifuge. Place the column in a clean 1.5 ml Lo-bind tube with the lid cut off.

32.

Add 25 ul of EB buffer to the column. Incubate 5 min at room temperature.

33.

Centrifuge 1 min at 13,000 rpm. Add the eluate back through the column to increase yield. Incubate for 5 min at room temperature. Centrifuge 1 min at 13,000 rpm.

34.

Transfer the eluate to a clean 0.5 ml Lo-bind tube.

35.

Dilute the library 1 in 20 in Ultrapure water (i.e., add 1 ul of the library to 19 ul of Ultrapure water). Vortex and pulse centrifuge.

36.

Libraries can be stored at -20 deg C until amplification. For long-term storage, store at -80 deg C.

Quant the library

37.

Make up the following master mix in a 1.5 ml Lo-bind tube. Vortex and pulse centrifuge.

A	B	C	D	E	F
Reagent	V2	C1	C2	V1	x _______ rxn
Ultrapure water	25 ul	na	na	15.9 ul
BSA	25 ul	10 mg /ml	0.4 mg/ml	1 ul
ABI Gold PCR Buffer	25 ul	10 X	1 X	2.5 ul
MgCl2	25 ul	25 mM	2.5 mM	2.5 ul
dNTPs	25 ul	25 mM	0.25 mM	0.25 ul
ABI Taq Gold DNA polymerase	25 ul	5 U/ul	0.05 U/ul	0.25 ul
SYBR Green	25 ul	5 X	0.12 X	0.6 ul
IS7	25 ul	10 uM	0.2 uM	0.5 ul
IS8	25 ul	10 uM	0.2 uM	0.5 ul

Assay A master mix

38.

Make up the following master mix in a 1.5 ml Lo-bind tube. Vortex and pulse centrifuge.

A	B	C	D	E	F
Reagent	V2	C1	C2	V1	x 16_ rxn
Ultrapure water	25 ul	na	na	15.9 ul	254.4 ul
BSA	25 ul	10 mg /ml	0.4 mg/ml	1 ul	16 ul
ABI Gold PCR Buffer	25 ul	10 X	1 X	2.5 ul	40 ul
MgCl2	25 ul	25 mM	2.5 mM	2.5 ul	40 ul
dNTPs	25 ul	25 mM	0.25 mM	0.25 ul	4 ul
ABI Taq Gold DNA polymerase	25 ul	5 U/ul	0.05 U/ul	0.25 ul	4 ul
SYBR Green	25 ul	5 X	0.12 X	0.6 ul	9.6 ul
CL107	25 ul	10 uM	0.2 uM	0.5 ul	8 ul
CL108	25 ul	10 uM	0.2 uM	0.5 ul	8 ul

Assay B master mix

39.

Add 24 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.

40.

Add 1 ul of DNA sample to the corresponding PCR tubes according to the scheme below. Pulse centrifuge the tubes.

A	B	C	D
PCR NTC	CL105_106_STD 10^3	dsLib001 Neat	dsLib005 Neat
PCR NTC	CL105_106_STD 10^3	dsLib001 1in20	dsLib005 1in20
CL105_106_STD 10^6	CL105_106_STD 10^2	dsLib002 Neat	...etc.
CL105_106_STD 10^6	CL105_106_STD 10^2	dsLib002 1in20
CL105_106_STD 10^5	dsCL104 +VE Neat	dsLib003 Neat
CL105_106_STD 10^5	dsCL104 +VE 1in20	dsLib003 1in20
CL105_106_STD 10^4	dsNTC -VE Neat	dsLib004 Neat
CL105_106_STD 10^4	dsNTC -VE 1in20	dsLib004 1in20

Assay A Plate set-up

A	B
0.1 uM CL104_duplex 1/500	CL105_106_STD 10^4
0.1 uM CL104_duplex 1/500	CL105_106_STD 10^4
PCR NTC	CL105_106_STD 10^3
PCR NTC	CL105_106_STD 10^3
CL105_106_STD 10^6	CL105_106_STD 10^2
CL105_106_STD 10^6	CL105_106_STD 10^2
CL105_106_STD 10^5
CL105_106_STD 10^5

Assay B Plate set-up

41.

Take the strip tubes to a post-PCR space. Place in thermal cycler and run the following program:

    95 deg C for 10 min



    Followed by 50 cycles of:

    

    95 deg C for 30 sec 

    60 deg C for 30 sec

    72 deg C for 30 sec

42.

Electrophorese 10 ul of the PCR product from the libraries (not standards) and controls on a 2% agarose gel.

2% Agarose Gel Electrophoresis

43.

Use the CT values from the qPCR to generate a standard curve for the standards in order to calculate how many template copies are present in each library. The positive control is used to calculate the efficiency of the library prep:

(# Copies of CL104 from Assay A / # copies CL104 from Assay B) * 100

Citation

Library preparation efficiency should typically be between 30-70% according to Gansauge and Meyer (2013). Molecule counts from the library preparation blank control should be less than 1x10^9, usually 1x10^8. The relationship between input volume of DNA extract and out of library molecules should be linear. If it is not, either too much DNA was used for library preparation or the DNA extract is inhibited. Gansauge et al. (2017) recommend to create a few preps with various input amounts to determine this; however, most of the time, this is not feasible because it is expensive to prepare multiple libraries for one sample. The Neat and 1in20 dilution of libraries should show be approximately 4.33 cycles apart. If they are not, there might be too much input DNA in the qPCR. Dilute futher and run the qPCR to get a more accurate estimate of library molecules. Insert sizes typically range from 20-120 bp. Indexing PCR will typically require 10-15 cycles of amplification.

This assay is also used to determine the number of cycles to give the indexing PCR, which needs to be stopped during the linear phase.

Index/amplify the library

44.

Make up the following master mix in a 1.5 ml Lo-bind tube. Ensure to prepare enough master mix for 4 reactions per library plus pipetting error. Vortex and pulse centrifuge.

Note

Remember that each library will have it's own unique combination of forward and reverse indexing primers. Do not add these to the master mix , but add each to each reaction individually! Take great care not to cross-contaminate primers: only have one tube open at a time. Use qPCR tubes with individual capped lids (not strip lids!).

Note

Ideally, indexing combinations should never be reused in the lab. Be sure to follow Illumina's recommendations when chosing primer combinations (e.g., ensure adequate diversity in the bases, ensure each is at least 3 bp different from each other, don't use indexes that will begin with two dark cycles, etc.). For instance, the NextSeq cannot read "GG" a the start of an index (so indexes should not end in "CC" as they are sequenced in the reverse complement).

A	B	C	D	E	F
Reagent	V2	C1	C2	V1	x _____ rxn
Ultrapure water	25 ul	na	na	10.9
BSA	25 ul	10 mg /ml	0.4 mg/ml	1 ul
ABI Gold PCR Buffer	25 ul	10 X	1 X	2.5 ul
MgCl2	25 ul	25 mM	2.5 mM	2.5 ul
dNTPs	25 ul	25 mM	0.25 mM	0.25 ul
ABI Taq Gold DNA polymerase	25 ul	5 U/ul	0.05 U/ul	0.25 ul
SYBR Green	25 ul	5 X	0.12 X	0.6 ul
P5_indexing_primer	25 ul	10 uM	0.2 uM	0.5 ul	Don't add to master mix
P7_indexing_primer	25 ul	10 uM	0.2 uM	0.5 ul	Don't add to master mix

45.

Add 19 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.

46.

Add 0.5 ul of the corresponding forward indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.

47.

Add 0.5 ul of the corresponding reverse indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.

48.

Add 5 ul of DNA sample to the corresponding reaction tubes according to the scheme below. Pulse centrifuge the tubes.

e.g.,

A	B	C
dsLib001	dsLib003	dsLib005
dsLib001	dsLib003	dsLib005
dsLib001	dsLib003	dsLib005
dsLib001	dsLib003	dsLib005
dsLib002	dsLib004	...etc.
dsLib002	dsLib004
dsLib002	dsLib004
dsLib002	dsLib004

49.

Take the strip tubes to a post-PCR space. Place in qPCR machine and run the following program:

    95 deg C for 10 min



 Followed by _________ cycles of:

    

    95 deg C for 30 sec 

    60 deg C for 30 sec

    72 deg C for 30 sec

Note

Note that the number of cycles to give should be determined based on the Assay A qPCR: stop while amplification is in the linear phase (before plateau). This PCR can be performed on a standard thermal cycler (and with your reagents of choice) but I prefer to run it on a qPCR as Assay A and B so I can monitor the amplification in real time. Ensure that you use a high-fidelity polymerase. You can use a proof-reading polymerase if doing standard PCR but do not use one with qPCR.

Purify the libraries

50.

Pulse centrifuge the PCR tubes. Combine replicate PCR reactions into a 1.5 ml Lo-bine Safelock tube. Vortex and pulse centrifuge.

51.

Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 1.6X beads : reaction volume (i.e., 160 ul). Follow the guidelines below:

http://www.gelifesciences.co.jp/catalog/pdf/SeraMagSelect_UserGuide.pdf

Elute in 35 ul of Ultrapure water.

Quantitate the libraries

52.

Dilute the libraries 1 in 10 in Ultrapure water (i.e., 1 ul library in 9 ul Ultrapure water).

53.

Use a LabChip GXII or equivalent fragment analyser (HiSense kit) to measure the molarity of the libraries between 160-500 bp.

http://www.perkinelmer.com/Content/LST_Software_Downloads/LabChip_GX_User_Manual.pdf

Citation

Libraries will be insert size + 136 bp, so the smallest fragments of interest will be ca. 166 bp (30 bp insert).

Pool libraries

54.

Pool libraries in equimolar concentrations such that the total amount of DNA per library does not exceed 500-1000 ng.

Note

If you are proceeding directly with hybridisation capture,STOP HERE and move to (e.g.) the protocol below . Try to pool libraries such that the total amount of DNA per library does not exceel 500 ng (the recommended input amount per capture).

hyRAD (Suchan et al. 2016; Grealy et al.)

55.

Use a Vivaspin 500 (MWCO 30,000 Da) centrifugal column to concentrate each library to 20-40 ul. Centrifuge at 15,000 rcf with the membrane facing outwards for 30 sec at a time.

http://www.sartorius.com/shop/ww/en/usd/applications-laboratory-filtration-ultrafiltration/vivaspin-500%2C-30%2C000-mwco-pes%2C-25pc/p/VS0121

Alternatively, concentrate the libraries using a SpeedyVac system, following the manufacturer's instructions.

http://assets.thermofisher.com/TFS-Assets/LED/manuals/DNA130%20User%20Manual%20final%20versionpdf.pdf

Size select and purify

56.

Run each pool in duplicate across two lanes (20 ul each) of a PippinHT electrophoresis system (2% gel, Marker 20B), selecting fragments between 160-500 bp and following the manufacturer's instructions:

http://www.sagescience.com/wp-content/uploads/2015/10/PippinHT-Operations-Manual-Rev-B_460005.pdf

57.

Combine replicates. Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 2X beads : reaction volume (i.e., 160 ul). Follow the guidelines below:

http://www.gelifesciences.co.jp/catalog/pdf/SeraMagSelect_UserGuide.pdf

Elute in 25 ul of Ultrapure water.

Quantitate the final library

58.

Dilute the libraries 1/2, 1/5, 1/10 in Ultrapure water (i.e., create a serial dilution in 10 ul volume).

59.

Measure the concentration of the neat library and these dilutions in duplicate on the Qubit following the manufacturer's instructions.

http://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0017209_Qubit_4_Fluorometer_UG.pdf

60.

Measure the molarity of the neat library and dilutions on a LabChip GXII Hisense kit (or equivalent fragment analyser) following the manufacturer's instructions:

http://www.perkinelmer.com/Content/LST_Software_Downloads/LabChip_GX_User_Manual.pdf

61.

Based on the average fragment length and Qubit measurement, calculate the molarity of the library dilutions. Create a standard curve to check that the concentrations are linear. If they can be "trusted", extrapolate the neat concentration based on the dilutions. Average all the measurements of the neat concentration to get the best estimate of the library molarity.

Sequencing

62.

Dilute the library to between 2-4 nM in Ultrapure water.

63.

Follow the manufacturer's instructions to perform the sequencnig run on your platform of choice.

DSBEST v2.0 (Caroe et al. 2018)

Disclaimer

Abstract

Before start

Attachments

Steps

Preparation

Prepare adapters

Determine the DNA input amount and concentration of adapters required

End Repair

Adapter ligation

Fill in

Purify library

Quant the library

Index/amplify the library

Purify the libraries

Quantitate the libraries

Pool libraries

Size select and purify

Quantitate the final library

Sequencing

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