Sep 25, 2023
DSBEST v2.0 (Caroe et al. 2018)
- 1Australian National University
Protocol Citation: Alicia Grealy 2023. DSBEST v2.0 (Caroe et al. 2018). protocols.io https://dx.doi.org/10.17504/protocols.io.j8nlkemwxl5r/v1
Manuscript citation:
Caroe C, Gopalakrishnan S, Vinner L, Mak SST, Sinding MHS, Samaniego JA, Wales N, Sicheritz-Ponten T, Gilbert MTP. 2018. Single-tube library preparation for degraded DNA. Methods in Ecology and Evolution, 9: 410-419.
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License,  which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 12, 2020
Last Modified: September 25, 2023
Protocol Integer ID: 34125
Disclaimer
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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!
Attachments
Caroe_et_al_2018_Sup...
10.7MB
Guidelines
Use at your own risk. You are responsible for double-checking that everything is correct!
Familiarise yourself with the relevent literature (Caroe et al. 2018) before attempting!
Materials
MATERIALS
Bst 2.0 WarmStart DNA Polymerase - 1,600 unitsNew England BiolabsCatalog #M0538S
T4 DNA Polymerase - 150 unitsNew England BiolabsCatalog #M0203S
T4 DNA Ligase - 20,000 unitsNew England BiolabsCatalog #M0202S
MinElute PCR Purification KitQiagenCatalog #28004
Tris-HCl, pH 8.0 (UltraPure)Thermo Fisher ScientificCatalog #15568025
5 M Sodium chloride (NaCl)Sigma AldrichCatalog #S5150-1L
DMSO SigmaCatalog #D8418
T4 Polynucleotide KinaseNew England BiolabsCatalog #M0201S
UltraPure™ DNase/RNase-Free Distilled WaterThermo FisherCatalog #10977049
AmpliTaq Gold™ DNA Polymerase with Gold Buffer and MgCl2Thermo FisherCatalog #4311806
SYBR™ Green I Nucleic Acid Gel Stain - 10,000X concentrate in DMSOThermo FisherCatalog #S7563
Sera-Mag Speed BeadsGe HealthcareCatalog #65152105050250
UltraPure 0.5M EDTA pH 8.0Invitrogen - Thermo FisherCatalog #15575020
Tween 20Sigma AldrichCatalog #P2287-100ml
Bovine Serum Albumin (BSA)Catalog #BSA-50
dNTPs 100 mM ea.BiolineCatalog #BIO-39025
T4 DNA ligase reaction bufferNew England BiolabsCatalog #Supplied with M0202S
Isothermal amplification bufferNew England BiolabsCatalog #Supplied with M0538S
PEG-4000 50% w/vNew England BiolabsCatalog #Supplied with M0202S
| Oligos | Concentration | Working concentration | Synthesis scale | Purification | Supplier | Resuspension buffer | Sequence 5'-3' | |
| IS1 | 500 uM | na | 100 nmol | HPLC | IDT | 10 mM Tris-HCl | A*C*A*C*TCTTTCCCTACACGACGCTCTTCCG*A*T*C*T | |
| IS2 | 500 uM | na | 100 nmol | HPLC | IDT | TE | G*T*G*A*CTGGAGTTCAGACGTGTGCTCTTCCG*A*T*C*T | |
| IS3 | 500 uM | na | 100 nmol | HPLC | IDT | TE | A*G*A*T*CGGAA*G*A*G*C/3SpC3/ | |
| CL104_duplex (order as double-stranded DNA) | 100 uM | 0.1 uM | 100 nmol | HPLC | IDT | TET buffer | /5Phos/TCGTCGTTTGGTATGGCTTCATTCAGCTCCGGTTCCCAACGATCAAGGCGAGTTACATGA/3Phos/ | |
| CL107 | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | TCATGTAACTCGCCTTGATCGT | |
| CL108 | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | TCGTCGTTTGGTATGGCTTC | |
| IS7 | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | ACACTCTTTCCCTACACGAC | |
| IS8 | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | GTGACTGGAGTTCAGACGTGT | |
| CL72_Custom_Seq_Primer | 100 uM | na | 25 nmol | desalted | IDT | Water | ACACTCTTTCCCTACACGACGCTCTTCC | |
| CL72_i5_index | 100 uM | na | 25 nmol | desalted | IDT | Water | GGA AGA GCG TCG TGT AGG GAA AGA GTG T | |
| CL105_CL106_Std | 100 uM | Dilute to 10^11 - 10^2 copies/ul | 4 nmol Ultramer | desalted | IDT | TET buffer | ACACTCTTTCCCTACACGACGCTCTTCCTCGTCGTTTGGTATGGCTTCTATCGUATCGATCGATCGACGATCAAGGCGAGTTACATGAAGATCGGAAGAGCACACGTCTGAACTCCAGTCAC | |
| P5-indexing primer | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | AATGATACGGCGACCACCGAGATCTACACNNNNNNNNACACTCTTTCCCTACACGACGCTCTT | |
| P7-indexing primer | 100 uM | 10 uM | 25 nmol | desalted | IDT | Water | CAAGCAGAAGACGGCATACGAGATNNNNNNNNGTGACTGGAGTTCAGACGTGT |
Table 1. Oligos needed. Store all oligos at -20 deg C.
Equipment needed
Thermalcycler
Bench-top centrifuge
QuantStudio 3 qPCR machine
P1000, P200, P100, P20, P10, P2 pipettes and extra-long filter tips
10% Household bleach solution
70% Ethanol
Kimwipes and paper towels
15 ml Falcon tubes
1.5 ml Safelock tubes
1.5 ml Lo-bind Safelock tubes
0.5 ml Lo-bind Safelock tubes
0.2 ml Lo-bind PCR tubes
8-well strip optical qPCR tubes with attached lid (0.1 ml profile)
Sharps container
UV glove box
96-well PCR plate rack
1.5 ml-2.0ml tube racks
15 ml tube racks
50 ml tube racks
Minispin
Vortex
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.
Preparation
Preparation
30m
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
Prepare adapters
40m
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.
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
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
Determine the DNA input amount and concentration of adapters required
30m
Measure the concentration of your DNA extract on the Qubit following the manufacturer's instructions.
If possible, run the DNA on a fragment analyser or gel electrophoresis to determine the fragment length distribution.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
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.
Make up 2 L of 1X TAE buffer:
50 ml of 40X TAE buffer
1950 ml of MilliQ water
Mix by inversion.
Using an electronic balance, weigh out 2.2 g of agarose powder on to a weigh boat using a spatula.
Note
Note that gel concentration can be adjusted. The more concentrated the gel, the greater the resolution of small fragment sizes.
Transfer the powder to a 200 ml conical flask.
Using the graduated measurng cylinder, measure out 110 ml of 1xTAE buffer. Add to the conical flask containing the agarose powder. Swirl the flask gently to mix.
Microwave the flask uncovered for 1 minute.
Safety information
A lid can loosly be placed over the flask but DO NOT tighten--allow steam to escape. Do not microwave for more than 1 minute at a time.
Do not fill flasks or beakers/Schott bottles more than half full with liquid!
Remove the flask from the microwave using oven mitts and swirl gently.
Safety information
The liquid is boiling! Use oven mitts to handle flask. Do not put your face over the opening to the flask as liquid can splash out!
Microwave the flask for a further minute but remove from the microwave if the agarose appears to boil excessively. Swirl to mix and examine near a light source to ensure the agarose has melted. Allow to cool for 5 minutes.
Place the gel casting tray into a rubber vice that will seal the ends tightly, or tape the ends with masking tape.
Place the assembly on a flat bench and use the spirit level to check it is level--adjust if needed.
Place a 20-well comb into the casting tray.
When the flask is cool to the touch, add 5 ul of SYBR Safe and swirl gently to mix. Avoid generating bubbles.
Safety information
Wait until the liquid is warm (not boiling) to cast the gel, or the tray may crack!
Pour the liquid gel slowly into the casting tray. Pop any bubbles that have formed using a clean pipette tip.
Let the gel set for 20-30 minutes at room temperature.
Allow residual gel to set in the flask, then scrape into the bin. Fill the flask half full with water and microwave until the water boils. Pour the water down the sink and clean the flask using a bottle brush.
When the gel is set, remove the combs gently.
Place the casting try and gel in the electrophoresis tank.
Fill the electrophoresis tank with 1 X TAE buffer to the fill line indicated on the tank.
Pipette 3 ul of 50 bp DNA ladder into the first well of the gel.
Note
The recommended volume will depend on the concentration of the ladder. Check the manufacterer's recommendations. If the ladder is not pre-mixed with loading dye, be sure to add 1-2 ul of loading dye before loading into the gel.
Place some Parafilm across a 96-well PCR plate rack and press down firmly to create small wells.
For each sample, pipette 1-2 ul of loading dye onto the Parafilm, taking care not to pierce the Parafilm.
Mix 10 ul of PCR product with the loading dye by pipetting gently up and down.
Transfer the 12 ul of PCR product/loading dye to the wells of the gel, taking care not to pierce the bottom of the well with the pipette tip.
Note
The volume each well can take will depend on the size of the comb used. Be sure not to overload the wells or product will float out the top of the well.
Place the lid on the gel tank and plug the electrodes into the appropriate power slots. Ensure the positive electrode is at the base of the gel.
Safety information
Take care when working with electricity and water!
Check electrical cords of all equipment and ensure none are damaged and that cords are not a tripping hazard. Do not use if the electrical cord is damaged in any way. Tag the instrument with warnings, make the area safe, and notify your line manager and anyone else in the immediate area that may be affected.
Use electrical equipment indoors only in an area free of explosive material, corrosive gas, powerful vibrations, direct exposure to sunlight, and temperature fluctuations. Use in a space where cables will not come into contact with liquids, be manually damaged, or interfere with other workplace operations.
Do not use electrical equipment with any other power adapter or cord than the one supplied.
Switch the power pack on a set the voltage to 80 V and the time to 1 hr and 10 min.
Note
Note that the voltage and time can be adjusted to suit what you are running on the gel. For amplicons (one small product), I will run the gel at 96 V for 30-40 min. For shotgun libraries, I will run the gel as above. The lower the voltage and longer it is run, the greater the separation of fragments will be.
Press 'Run' or 'Start' on the power pack and check to see that bubbles are rising from electrodes.
When the run is over, switch of the power pack, remove the lid, and remove the gel from the tank, taking care not to let it slide off the tray.
Safety information
Do not remove the lid to the electrophoresis tank until the power pack is switched off.
Place the gel on the UV transilluminator and photograph using the attached camera. Follow the manufactuerer's instructions to use the equipment.
Safety information
Take care working with UV. You should have UV safety training. Do not open the transilluminator while the UV is on! Use signage to warn others when the UV is on.
Discard the gel into a designated biohazard bin, and clean the UV dock with 70% ethanol.
Dispose of used tips into a designated sharps container.
Dispose of gel waste into a biohazard bag.
Used combs, beakers, flasks, and tray should be washed with warm water and placed on a rack to dry.
Gloves and chemical waste should be sealed in a biohazard bag for incineration.
Calculate the molarity of the dsDNA ends. Use the calculator at:
Expected result
e.g., 1 ul of 0.1 uM CL104_duplex
Mass dsDNA input (ng) = 3.71 ng
Length (bp) = 60 bp
pmol dsDNA ends = 2*(((ng-input/1000/1000)/((length*617.96)+36.04))*1000*1000*1000)
= 0.2 pmol
Input ds copy number = ((pmol dsDNA ends/2)/1000/1000/1000/1000)*(6.022E+23)
= 6.02E+10
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.
Expected result
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
End Repair
1h 10m
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 |
Aliquot 8 ul into a 0.2 ml Lo-bind PCR tube for each reaction.
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.
Incubate in a thermal cycler at:
20 deg C for 30 min
65 deg C for 30 min
4 deg C hold
Adapter ligation
Adapter ligation
50m
Add 1 ul of the DS_adapter_mix DILUTION (e.g., 10 uM) to the finished end-repair reaction above. Vortex and pulse centrifuge.
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 |
Add 9 ul to each reaction. Vortex and pulse centrifuge.
Incubate in a thermal cycler at:
20 deg C for 30 min
65 deg C for 10 min
4 deg C hold
Fill in
Fill in
40m
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 |
Add 10 ul of the above mix to each reaction. Vortex and pulse centrifuge.
Incubate in a thermal cycler at:
65 deg C for 15 min
80 deg C for 15 min
4 deg C hold
Purify library
Purify library
30m
Purify the libraries using a QIAGEN MinElute PCR Purification kit.
Note
Be sure to store the silica columns at 4 deg C.
Briefly...
Combine 300 ul of PB Buffer with each reaction in a 1.5 ml Lo-Bind tube. Vortex and pulse centrifuge.
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.
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.
Repeat Step 29.
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.
Add 25 ul of EB buffer to the column. Incubate 5 min at room temperature.
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.
Transfer the eluate to a clean 0.5 ml Lo-bind tube.
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.
Libraries can be stored at -20 deg C until amplification. For long-term storage, store at -80 deg C.
Quant the library
Quant the library
2h 30m
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
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
Add 24 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.
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
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
Electrophorese 10 ul of the PCR product from the libraries (not standards) and controls on a 2% agarose gel.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Make up 2 L of 1X TAE buffer:
50 ml of 40X TAE buffer
1950 ml of MilliQ water
Mix by inversion.
Using an electronic balance, weigh out 2.2 g of agarose powder on to a weigh boat using a spatula.
Note
Note that gel concentration can be adjusted. The more concentrated the gel, the greater the resolution of small fragment sizes.
Transfer the powder to a 200 ml conical flask.
Using the graduated measurng cylinder, measure out 110 ml of 1xTAE buffer. Add to the conical flask containing the agarose powder. Swirl the flask gently to mix.
Microwave the flask uncovered for 1 minute.
Safety information
A lid can loosly be placed over the flask but DO NOT tighten--allow steam to escape. Do not microwave for more than 1 minute at a time.
Do not fill flasks or beakers/Schott bottles more than half full with liquid!
Remove the flask from the microwave using oven mitts and swirl gently.
Safety information
The liquid is boiling! Use oven mitts to handle flask. Do not put your face over the opening to the flask as liquid can splash out!
Microwave the flask for a further minute but remove from the microwave if the agarose appears to boil excessively. Swirl to mix and examine near a light source to ensure the agarose has melted. Allow to cool for 5 minutes.
Place the gel casting tray into a rubber vice that will seal the ends tightly, or tape the ends with masking tape.
Place the assembly on a flat bench and use the spirit level to check it is level--adjust if needed.
Place a 20-well comb into the casting tray.
When the flask is cool to the touch, add 5 ul of SYBR Safe and swirl gently to mix. Avoid generating bubbles.
Safety information
Wait until the liquid is warm (not boiling) to cast the gel, or the tray may crack!
Pour the liquid gel slowly into the casting tray. Pop any bubbles that have formed using a clean pipette tip.
Let the gel set for 20-30 minutes at room temperature.
Allow residual gel to set in the flask, then scrape into the bin. Fill the flask half full with water and microwave until the water boils. Pour the water down the sink and clean the flask using a bottle brush.
When the gel is set, remove the combs gently.
Place the casting try and gel in the electrophoresis tank.
Fill the electrophoresis tank with 1 X TAE buffer to the fill line indicated on the tank.
Pipette 3 ul of 50 bp DNA ladder into the first well of the gel.
Note
The recommended volume will depend on the concentration of the ladder. Check the manufacterer's recommendations. If the ladder is not pre-mixed with loading dye, be sure to add 1-2 ul of loading dye before loading into the gel.
Place some Parafilm across a 96-well PCR plate rack and press down firmly to create small wells.
For each sample, pipette 1-2 ul of loading dye onto the Parafilm, taking care not to pierce the Parafilm.
Mix 10 ul of PCR product with the loading dye by pipetting gently up and down.
Transfer the 12 ul of PCR product/loading dye to the wells of the gel, taking care not to pierce the bottom of the well with the pipette tip.
Note
The volume each well can take will depend on the size of the comb used. Be sure not to overload the wells or product will float out the top of the well.
Place the lid on the gel tank and plug the electrodes into the appropriate power slots. Ensure the positive electrode is at the base of the gel.
Safety information
Take care when working with electricity and water!
Check electrical cords of all equipment and ensure none are damaged and that cords are not a tripping hazard. Do not use if the electrical cord is damaged in any way. Tag the instrument with warnings, make the area safe, and notify your line manager and anyone else in the immediate area that may be affected.
Use electrical equipment indoors only in an area free of explosive material, corrosive gas, powerful vibrations, direct exposure to sunlight, and temperature fluctuations. Use in a space where cables will not come into contact with liquids, be manually damaged, or interfere with other workplace operations.
Do not use electrical equipment with any other power adapter or cord than the one supplied.
Switch the power pack on a set the voltage to 80 V and the time to 1 hr and 10 min.
Note
Note that the voltage and time can be adjusted to suit what you are running on the gel. For amplicons (one small product), I will run the gel at 96 V for 30-40 min. For shotgun libraries, I will run the gel as above. The lower the voltage and longer it is run, the greater the separation of fragments will be.
Press 'Run' or 'Start' on the power pack and check to see that bubbles are rising from electrodes.
When the run is over, switch of the power pack, remove the lid, and remove the gel from the tank, taking care not to let it slide off the tray.
Safety information
Do not remove the lid to the electrophoresis tank until the power pack is switched off.
Place the gel on the UV transilluminator and photograph using the attached camera. Follow the manufactuerer's instructions to use the equipment.
Safety information
Take care working with UV. You should have UV safety training. Do not open the transilluminator while the UV is on! Use signage to warn others when the UV is on.
Discard the gel into a designated biohazard bin, and clean the UV dock with 70% ethanol.
Dispose of used tips into a designated sharps container.
Dispose of gel waste into a biohazard bag.
Used combs, beakers, flasks, and tray should be washed with warm water and placed on a rack to dry.
Gloves and chemical waste should be sealed in a biohazard bag for incineration.
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
Expected result
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
Index/amplify the library
2h 30m
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 |
Add 19 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.
Add 0.5 ul of the corresponding forward indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.
Add 0.5 ul of the corresponding reverse indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.
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 | D | E | F | |
| dsLib001 | dsLib003 | dsLib005 | ||||
| dsLib001 | dsLib003 | dsLib005 | ||||
| dsLib001 | dsLib003 | dsLib005 | ||||
| dsLib001 | dsLib003 | dsLib005 | ||||
| dsLib002 | dsLib004 | ...etc. | ||||
| dsLib002 | dsLib004 | |||||
| dsLib002 | dsLib004 | |||||
| dsLib002 | dsLib004 |
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
Purify the libraries
Pulse centrifuge the PCR tubes. Combine replicate PCR reactions into a 1.5 ml Lo-bine Safelock tube. Vortex and pulse centrifuge.
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:
Elute in 35 ul of Ultrapure water.
Quantitate the libraries
Quantitate the libraries
Dilute the libraries 1 in 10 in Ultrapure water (i.e., 1 ul library in 9 ul Ultrapure water).
Use a LabChip GXII or equivalent fragment analyser (HiSense kit) to measure the molarity of the libraries between 160-500 bp.
Expected result
Libraries will be insert size + 136 bp, so the smallest fragments of interest will be ca. 166 bp (30 bp insert).
Pool libraries
Pool libraries
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).
Protocol
NAME
hyRAD (Suchan et al. 2016; Grealy et al.) CREATED BY
Alicia Grealy
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.
| Tube | Qty | For ... | |
| 1.5 ml Lo-bind Safelock tube | 1 | 10 mM Tris-HCl | |
| 1.5 ml Lo-bind Safelock tube | 1 | 10X Annealing Buffer | |
| 0.5 ml Lo-bind Safelock tube | 2 | P1 and P2 adapter oligos | |
| 0.2 ml Lo-bind PCR tube | 1 | Annealing P1 and P2 adapter oligos |
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.
| Buffer | Reagent | Volume to add | Final concentration in solution | |
| 10 mM Tris-HCl | 1 M Tris-HCl | 10 ul | 10 mM | |
| Ultrapure water | 990 ul | na | ||
| 10 X Annealing Buffer | 1 M Tris-HCl | 100 ul | 100 mM | |
| 5 M NaCl | 100 ul | 500 mM | ||
| 0.5 M EDTA | 20 ul | 10 mM | ||
| Ultrapure water | 780 ul | na |
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!
| Working stock | Reagent | Volume to add | |
| 10 uM P5_Indexing_Primer | 100 uM Stock | 50 ul | |
| Ultrapure water | 450 ul | ||
| 10 uM P7_Indexing_Primer | 100 uM Stock | 50 ul | |
| Ultrapure water | 450 ul |
Pre-program the thermal cycler.
Combine the following in a 0.2 ml Lo-bind PCR tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | |
| P1.1 adapter oligo (EcoRI) | 100 ul | 100 uM | 10 uM | 10 ul | |
| P1.2 adapter oligo (EcoRI) | 100 ul | 100 uM | 10 uM | 10 ul | |
| Annealing buffer | 100 ul | 10 X | 1 X | 10 ul | |
| Ultrapure water | 100 ul | na | na | 70 ul |
Combine the following in a 0.2 ml Lo-bind PCR tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | |
| P2.1 adapter oligo (MspI) | 100 ul | 100 uM | 10 uM | 10 ul | |
| P2.2 adapter oligo (MspI) | 100 ul | 100 uM | 10 uM | 10 ul | |
| Annealing buffer | 100 ul | 10 X | 1 X | 10 ul | |
| Ultrapure water | 100 ul | na | na | 70 ul |
Incubate in a thermal cycler at:
95 deg C for 1 min
Cool at a rate of 0.1 deg C / sec until the solution reaches 20 deg C
Store at -20 deg C.
Enzymatic digestion
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 2.2 Master mix | |
| 0.2 ml Lo-Bind PCR tube | # of samples | Enzymatic digestion for samples |
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | |
| CutSmart Buffer (NEB) | 50 ul | 10 X | 1 X | 5 ul | |
| EcoRI-HF (NEB) | 50 ul | 20 U/ul | 0.4 U/ul (20 U) | 1 ul | |
| MspI (NEB) | 50 ul | 20 U/ul | 0.4 U/ul (20 U) | 1 ul | |
| Ultrapure water | 50 ul | na | na | 33 ul |
Note
EcoRI-HF (NEB) is active for >8 hr.
MspI (NEB) is active between 2-4 hr.
Note: This reaction has been scaled up to a 50 ul reaction as NEB recommends not leaving 10 ul reactions longer than 1 hr due to evaporation. Also, 10-20 U fo enzyme is recommended to digest 1 ug of DNA in a reaction volume of 50 ul for 1 hr. So, to ensure complete digestion of 1 ug of genomic DNA, 20 U of each enzyme is used n a reaction volume of 50 ul for 4 hr. MspI is not active beyond 4 hr. Also, the enzyme volume should not exceed 10% of the total reaction volume to prevent star activity due to excess glycerol.
Consider performing several reactions to obtain enough DNA for quality control along the way. I have started with 4 ug of DNA (split into 4 reactions--two DNA extracts performed in duplicate).
The ddRAD protocol recommends NOT to heat-denature restriction enzymes as they will be removed ruing the SPRI bead cleanup that follows below.
Add 40 ul of master mix to 10 ul of DNA (at 100 ng/ul ca. 1000 ng total) in a 0.2 ml Lo-bind PCR tube. Vortex and pulse centrifuge.
Note
Use high-quality DNA: quantify the concentration using the Qubit 3.0 fluorometer BR kit following the manufacturer's instructions:
Estimate the purity of the DNA using a NanoDrop spectrophotometer, following the manufacturer's instructions:
Note: do not pay attention to the DNA quanitity proveided by the NanoDrop or fragment analyser--the Qubit is much more accurate as it measures double-stranded DNA only.
Run the DNA on a fragment analyser or gel electrophoresis to determine the fragment length distribution:
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Expected result
DNA needs to be 100 ng/ul in 10 ul volume (i.e., 1 ug). Dilute to the sample to this concentration in Ultrapure water if needed.
e.g., I typically begin with samples that measure 1100 ng/ul in 100 ul (110 ug total):
In the example below, sample MD#033 was 1000 ng/ul in 100 ul
MD#034 was 1100 ng/ul in 100 ul
Pure DNA should have a 260/280 ratio of between 1.8-2.0 and a 230/260 ratio of 2.0-2.2. If the DNA is not pure, consider cleaning the neat extract using your method of choice (e.g., sodium-acetate/ethanol precipitation, etc.)
Fresh tissue should yield a high molecular weight band on an agarose gel (i.e., above 10 kb), with minimal smearing (as smearing indicates degradation).
Figure 1. An example of decently high-quality DNA extracts run on a 2% agarose gel electrophoresis (@ 80V for 1 hr 10 min), though there is some degradation.
Note
Add the DNA in a physically separate space, suitable for 'modern' DNA (i.e., NOT inside an ultraclean environment).
Incubate in a thermal cycler at:
37 deg C for 4 hr
Hold at 4 deg C
Combine any replicates.
SPRI bead clean-up
Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 2X beads : reaction volume (i.e., 160 ul). Follow the guidelines below:
https://www.gelifesciences.co.jp/catalog/pdf/SeraMagSelect_UserGuide.pdf
Elute in 20 ul of 10 mM Tris-HCl.
Note
This step is to remove the enzymes only. To avoid losing product, use a ration of 1.8-2X beads, which should keep everything above 100 bp without too much yield loss.
Qubit
Measure the concentration using the Qubit HS kit following the manufacturer's instructions.
Expected result
Expect a loss of around 3%.
e.g., MD#034 = 97 ng/ul in 20 ul (or 1940 ng total). 2 ug of this sample was input into the reaction, so the loss is approximately 3%.
Agarose gel electrophoresis
Electrophorese 2 ul of product (200-500 ng in 5-10 ul) on a 2% agarose gel.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Expected result
Digested DNA should show a smear rather than a large band. However, some high-molecular weight fragments may remain but they won't be the same size as previously--it's just difficult to resolve these fragments on a gel (i.e., it is difficult to distinguish between 30,000 bp and 10,000 bp on this kind of gel).
Note: I have tested putting more enzyme into the reaction the results look the same.
Figure 2. An example of DNA that has undergone the restriction digest with EcoRI-HF and MspI using the reaction conditions described above.
Adapter ligation
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 3.2 Master mix | |
| 0.2 ml Lo-Bind PCR tube | # of samples x 2 | Reactions, two per sample |
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | |
| CutSmart Buffer (NEB) | 20 ul | 10 X | 1 X | 2 ul | |
| P1 adapter | 20 ul | 10 uM | 0.5 uM | 1 ul | |
| P2 adapter | 20 ul | 20 uM | 1 uM | 1 ul | |
| ATP | 20 ul | 100 mM | 1 mM | 0.2 ul | |
| T4 DNA ligase | 20 ul | 400 U/ul | 20 U/ul (400 U) | 1 ul | |
| Ultrapure water | 20 ul | na | na | 5.8 ul |
Note
Peterson et al. suggests to use 2-10 fold adapters : sticky ends in a 40 ul reaction volume. Based on the approximate average fragment size and the # ng input of DNA, calculate the molarity of the DNA ends using:
e.g., 500 ng of 275 bp DNA has 5.883 pmol DNA ends. So, for 10X adapters we would need ca. 30 pmol DNA ends per adapter, which, at ~45 bp, would be about 400 ng.
Molecular weight of P1 adapter = 32.7507 kDa. 10 uM P1 = 327.507 ng/ul
Molecular weight of P2 adapter = 18.3674 kDa. 20 uM P2 = 367.348 ng/ul
So using 1 ul of each at these concentrations should be enough for this quantity of input DNA.
e.g., if we have 14 ul of digested DNA at 97 ng/ul and we perform the ligation in duplicate, that would make the input 679 ng. If we estimate the average fragment length of the smear at 800 bp, this would be 2.7 pmol DNA ends. Even if we overestimated the fragment length (and it is actually 400 bp), the above amounts of adapter would still be in excess.
Adapter dimer will be removed with the size selection so it shouldn't be too bad if the adapters are in excess.
Add 11 ul of the above master mix to 9 ul of digested DNA in a 0.2 ml Lo-bind PCR tube. Vortex and pulse centrifuge.
Incubate in a thermal cycler at:
16 deg C for ca. 20 hr (16 hr - overnight)
Combine any replicates.
Note
NEB suggest to perform the ligation overnight.
SPRI bead clean-up
Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 2X beads : reaction volume (i.e., 160 ul). Follow the guidelines below:
https://www.gelifesciences.co.jp/catalog/pdf/SeraMagSelect_UserGuide.pdf
Elute in 21 ul of 10 mM Tris-HCl.
Note
This step is to remove the T4 DNA ligase, adapter dimer (ca. 90 bp) and unligated adapter (55 and 35 bp).
At this stage we "could" remove some of the fragments that are not of interest. A right-sided selection followed by a left-sided selection can be sued to select fragments within a certain range. Potentially a simple right-sided selection may be beneficial here because it will remove larger fragments without such a loss of yield of smaller fragments that may be of interest. Trying to remove small fragments of litte interest (e.g. <100 bp) may result in a substantial loss of yield for those in the target range (180-300 bp).
For a right-sided selection, a ratio of 0.6 X beads should remove most fragments >500 bp without substantial reduction in yield in the 100-300 bp size range.
For a left sided-selection, a ratio of 1.6 X beads should remove most fragments <100 bp without a substantial reduction in yield in the 100-300 bp size range.
BUT this would only really be beneficial if doing the amplification BEFORE size-selection.
Qubit
Measure the concentration using the Qubit HS kit following the manufacturer's instructions.
Expected result
Expect approximately 30% loss of yield.
e.g., MD#034 = 37.8 ng/ul (or 945 ng total), which is a 30% loss.
qPCR quant the RAD library
Note
This step is to make sure the ligation worked and to see what sized fragments would amplify. This step could be performed with alongside standards of known concentration to calculate the number of libarary molecules output in the RAD library.
You can also run more of a serial dilution of the library to ensure the quantitation is accurate. Here, I only input 1 ul of the neat library and 1 ul of a 1 in 20 dilution.
It could be possible to amplify the RAD library with indexes and THEN perform size selection. However, I am not sure how this may bias the probe set as it has not been tested, so I follow recommendations of Suchan et al. (2016), performing the size selection BEFORE the indexing PCR/library amplification.
After initial testing, I routinely skip this step
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 4.2 Master mix | |
| 8-well strip qPCR tubes 0.1 ul profile | 1 | PCR amplification |
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
| 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 |
Add 24 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.
Dilute each library 1 in 20 (i.e., 1 ul of library in 19 ul of Ultrapure water). Vortex and pulse centrifuge.
Add 1 ul of DNA sample to the corresponding PCR tubes according to the scheme below. Pulse centrifuge the tubes.
e.g.,
| PCR NTC | ||||
| PCR NTC | ||||
| MD#033 Neat | ||||
| MD#033 1 in 20 | ||||
| MD#034 Neat | ||||
| MD#034 1 in 20 | ||||
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
Agarose gel electrophoresis
Electrophorese 2 ul of product (200-500 ng in 5-10 ul) on a 2% agarose gel.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Expected result
The longest adapter dimer will be 112 bp but most should have been removed with the SPRI bead purification. Inserts of 50 bp will be 162 bp. If there are many fragments outside the target range (180 bp insert, i.e., 272 bp with adapters or say 250-300 bp), then definitely size select BEFORE indexing/amplification. If most amplified fragments are within the desired size range, it MAY be possible to size select after indexing/library amplification, but I have not tested this, and do not know whether it would bias the probe set in a different way. It is possible that size selecting after indexing/library amplification would deplete the yield so much as to require more amplification anyway.
Figure 3. An example of the size of fragments that would be amplified from the RAD library BEFORE size selection. It does appear that ligation worked. Amplification of the neat extract does appears to be inhibited, so if quantitating the number of molecules via qPCR using a standard, perhaps run a small dilution series for each library.
Size selection using Pippin HT
Note
The maximum input into the Pippin HT is 1 ug /lane sheared genomic DNA (i.e. 75 ng/ul in 20 ul)
The minimum input into the Pippin HT is 15 ng /lane sheared genomic DNA
Depending on the total yield and concentration determined using the Qubit (see Step 3.6 above), run approximately 1 ug of DNA in 20 ul (ca. 75 ng/ul) of each samples across a lane of a PippinHT electrophoresis system (2% gel, Marker 20B), selecting fragments 272 bp in size (272 bp peak with tight range 212-332 bp--this equates to an insert size of 180 bp) and following the manufacturer's instructions:
Note
180 bp insert size / probe size was used by Suchan et al. (2016) and Schmitt et al. (2018?). It is possible to make probes smaller than this, which may be beneficial for caputuring ancient DNA, but be sure to check the restriction enzymes used will generate enough loci within this size range.
SPRI cleanup
Combine any replicates. Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 2X beads : reaction volume. Follow the guidelines below:
Elute in 21 ul of Ultrapure water.
Note
This cleanup is for the purpose of buffer exchange, but also will concentrate any replicates into a smaller working volume.
Qubit
Measure the concentration using the Qubit HS kit following the manufacturer's instructions.
Expected result
Expect approximately 99% loss of yield.
e.g., 996 ng was input into the PippinHT lane for MD#033 and 9.724 ng came out (1%)
qPCR quant the size-selected RAD library
Note
This step is to make sure the size-selection worked. This qPCR could be performed alongside standards of known concentration to quantify the number of library molecules present in the RAD library after size selection.
After initial testing, I routinely skip this step.
Follow Steps 4.1 - 4.7 above to perform the qPCR if so desired.
Expected result
If the average insert size is 180 bp, then there should be a tight smear around ...
Index / amplify the RAD library
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 7.2 Master mix | |
| 8-well strip qPCR tubes 0.1 ul profile | 1 | PCR amplification |
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge. Ensure to prepare enough master mix for multiple reactions per library plus pipetting error.
Safety information
If you are not interested in sequencing the probes at all, then you can replace the indexing primers with IS7 and IS8 primers that will just amplify the library. If you aren't interested in knowing which probes came from which extract, then all reactions may be amplified with the same indexing primers.
Note
I have typically performed the indexing / library amplification in quadruplicate reactions, using 5 ul of library per reaction (using the up whole library). You can input less DNA into each reaction and perform more replicates if you desire more DNA at the end. This will mean you have more clonal probes, but that is not too much of an issue here because we do want several copies of each probe. You can also perform more cycles, though having little DNA and increasing the number of cycles can generate amplification artefacts. Typically I will perform 30 cycles, just enough to take the reaction to plateau.
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).
| Reagent | V2 | C1 | C2 | V1 | x _____ rxn | |
| Ultrapure water | 50 ul | na | na | 29.5 ul | ||
| KAPA High Fidelity Buffer | 50 ul | 5 X | 1 X | 10 ul | ||
| P5_indexing_primer | 50 ul | 5 uM | 0.2 uM | 2 ul | Do not add to master mix | |
| P7_indexing_primer | 50 ul | 5 uM | 0.2 uM | 2 ul | Do not add to master mix | |
| KAPA HiFi Hot Start DNA Polymerase | 50 ul | 1 U/ul | 0.02 U/ul | 1 ul | ||
| dNTPs | 50 ul | 25 uM | 0.25 uM | 0.5 ul |
Add 41 ul of master mix to the corresponding PCR tubes. Pulse centrifuge the tubes.
Add 2 ul of the corresponding forward indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.
Add 2 ul of the corresponding forward indexing primer to the appropriate reaction tube. Pulse centrifuge the tubes.
Add 5 ul of purified size-selected RAD library to the corresponding PCR tubes according to the scheme below. Pulse centrifuge the tubes.
Note
The number of ng of library input into each reaction for me is typically 1.25 - 2.21 ng. This is perhaps too little DNA for the number of cycles given, and may have generated an artefact. Potentially aim for 5-10 ng if possible.
e.g.,
| MD#033 | ||||
| MD#033 | ||||
| MD#033 | ||||
| MD#033 | ||||
| MD#034 | ||||
| MD#034 | ||||
| MD#034 | ||||
| MD#034 |
Take the strip tubes to a post-PCR space. Place in thermal cycler and run the following program:
98 deg C for 10 min
Followed by 30 cycles of:
985 deg C for 20 sec
60 deg C for 30 sec
72 deg C for 40 sec
Then a final extension of:
72 deg C for 10 min
Pool replicate reactions into a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
SPRI bead clean-up
Purify the libraries using SeraMag Speed Beads or SeraMag Select using a1.4X beads : reaction volume. Follow the guidelines below:
Elute in 40 ul of Ultrapure water.
Note
This cleanup is to remove PCR reagents and primer dimer, and to concentrate the replicates. Dimer should be around 160 bp. Fragments of interest are approximately 340 bp. This ratio of beads should remove almost everything below 200 bp.
Qubit
Measure the concentration using the Qubit HS kit following the manufacturer's instructions.
Expected result
Expect amplification to increase the DNA about 250-300 X.
e.g., Approximately 9 ng of MD#033 input across 4 PCR reactions turned into 2392 ng total, so yield increased 265-fold.
Approximately 5 ng of MD#034 input across 4 PCR reactions turned into 1520 ng total, so yield increased 304-fold.
Agarose gel electrophoresis
Electrophorese 5 ul of product (200-500 ng in 5-10 ul) on a 2% agarose gel.
Note
For any gel steps, a fragment analyser platform may alternatively be used, but I have found it to be much more straight forward to perform a gel.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Expected result
There should be a tight smear around a peak of 340 bp with little product below 200 bp. Insert sizes of 0 bp will be at 160 bp. Indexing dimer should be at 115 bp, but the purification should have removed most of this.
e.g.,
Figure 4. A large band can be seen around 350 bp, however, there is also a larger fragment above 450 bp present. This may be PCR artefact from too many cycles given a small input of DNA (see Belt and Demarini, 1991). This was not investigated further.
Use a LabChip GXII or equivalent fragment analyser (HiSense kit) to measure the molarity of the libraries.
Alternatively, use the Qubit concentration and the average fragment size seen on the gel to estimate the molarity of the libraries:
Pool amplified libraries
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Pooled RAD library | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Aliquot of RAD library for sequencing |
Pool libraries in equimolar concentrations such that the total volume is is approximately 100 ul.
Expected result
e.g., Combine 35 ul of MD#034 with 31.01 ul of MD#033. The total ng will be 3184.4 ng, the concentration will be 48.24 ng/ul.
Aliquot 20 ul of the pooled RAD library into a new 1.5 ml Lo-bind Safelock tube for future sequencing. Store at -20 deg C. See Steps 58-63 in the following protocol to quantitate and sequence the final RAD library.
Adapter removal
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 9.2 master mix | |
| 0.2 ml Lo-Bind PCR tube | 1 per 1 ug DNA | Adapter removal reaction |
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
Note
Perform enough replicate reactions for each 1 ug of DNA.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | x ______ rxn | |
| CutSmart Buffer (NEB) | 50 ul | 10 X | 1 X | 5 ul | ||
| MspI | 50 ul | 20 U/ul | 0.4 U/ul (20 U) | 1 ul | ||
| Ultrapure water | 50 ul | na | na | 25.4 ul |
Aliquot 31.4 ul of the above master mix into 0.2 ml Lo-bind PCR tubes.
Add 18.6 ul (1 ug) of pooled purified RAD library to each tube. Vortex and pulse centrifuge.
Note
Remember to adjust the water volume in the reactions to accomodate the volume of library added to the reaction.
Incubate the reactions in a thermal cycler at:
37 deg C for 4 hr
Combine the replicates.
SPRI bead clean-up
Purify the libraries using SeraMag Speed Beads or SeraMag Select using a 2X beads : reaction volume. Follow the guidelines below:
Elute in 30 ul of Ultrapure water.
Note
This cleanup is simply to remove the cut-off adapters (68 bp).
Qubit
Measure the concentration using the Qubit HS kit following the manufacturer's instructions.
Expected result
Expect a loss of about 56%.
e.g., After adapter removal, MD#033/MD#034 RAD library was 59.8 ng/ul or 1794 ng total. Before we had 3184 ng--so about 56% was lost.
Agarose gel electrophoresis
Electrophorese 5 ul of product (200-500 ng in 5-10 ul) on a 2% agarose gel.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
Expected result
50-53 bp are being cut-off during the adapter removal process, so the peak should be between 296-299 bp.
e.g.,
Figure 5. The RAD library after adapter removal. It is the right size.
In-vitro transcription and biotinylation
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Step 10.3 master mix | |
| 0.2 ml Lo-Bind PCR tube | # reactions | Transcription/biotinylation reaction | |
| 0.2 ml Lo-Bind PCR tube | 2 | Aliquots of TURBO DNase and SuperaseIn RNAse inhibitor | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Combine probes for cleanup | |
| RNeasy mini spin column | 1 | Purification | |
| 1.5 ml Lo-Bind Safelock tube with the lid cut off | 1 | Elution | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Final tube for probes | |
| 0.5 ml Lo-Bind Safelock tube | Depends on total ug of probes | 6-ul aliquots of probes |
Aliquot 10 ul of TURBO DNAse and 10 ul of SuperaseIn RNAse inhibitor into separate 0.2 ml Lo-bind tubes.
Combine the following in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
Note
Perform enough replicate reactions to transcribe all the library left into probes (inputting 500 ng of library per reaction).
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | x ______ rxn | |
| Reaction Buffer (HiScribe kit) | 20 ul | 10 X | 0.75 X | 1.5 ul | ||
| dATP | 20 ul | 100 uM | 7.5 uM | 1.5 ul | ||
| dCTP | 20 ul | 100 uM | 7.5 uM | 1.5 ul | ||
| dGTP | 20 ul | 100 uM | 7.5 uM | 1.5 ul | ||
| dUTP | 20 ul | 100 uM | 5 uM | 1 ul | ||
| biotin-UTP | 20 ul | 10 mM | 2.5 mM | 5 ul | ||
| T7 RNA polymerase mix | 20 ul | ? | ? | 1.5 ul | ||
| Ultrapure water | 20 ul | na | na | Adjust depending on the volume of library added |
Note
Typically I do not add water to the reaction.
Aliquot 13.5 ul of the above mater mix to 0.2 ml Lo-bind PCR tubes. Bring all tubes to a post-PCR space, including teh aliquots made in Step 10.2.
Concentrate the "probe set" (i.e., the pool RAD library with the adapters cut off) such that roughly 500 ng in 6.5 ul can be added per reaction using a SpeedVac, following the manufacturer's instructions:
Expected result
e.g., After quanting and the gel, I had 1495 ng of MD#033/MD#034 in 25 ul. I concentrated this to 16.5 ul (i.e., 90.6 ng/ul). Then I added 6.5 ul into two replicate transcription/biotinylation reactions (i.e., 589 ng was input into dupllicate reactions).
Add 6.5 ul of the "probe set" (ca. 500 ng) to each reaction. Vortex and pulse centrifuge.
Incubate in a thermalcycler at:
37 deg C for 16 hr
In the last 30 min, add 2 ul of TURBO DNase (2 U/ul, or 4 U for up to 10 ug). Pipette up and down to mix.
Combine replicate reactions in a 1.5 ml Lo-bind tube. Top up to 100 ul with RNase-free water.
Cleanup using an RNeasy Mini Kit
Note
This step is to remove reagents.
Follow the manufacturer's instructions to purify the probes using an RNeasy Mini Kit:
Elute in 60 ul RNase-free water.
Add 2.5 ul of SUPERase-IN RNAse Inhibitor (20 U/ul) to the eluate. Flick to mix and pulse centrifuge.
Qubit
Measure the concentration of the probes using the Qubit RNA kit following the manufacturer's instructions.
Expected result
The HiScribe kit suggests that 1 ug DNA --> 10 ug, but after inputting 1.2 ug DNA I got ~40 ug:
e.g., Measuring the concentration of a 1 in 2 and a 1 in 10 dilution of the probes (MD#033/MD#034) I estimate the neat to be either 23.7 or 55.8 ug of RNA, respectively. Averaging those values gives approximately 39.75 ug of RNA, or enough for about 40 hybridisation capture reactions (giving each reaction 1 ug of probes).
Fragment analyse
Examine the fragment length distribution of the probes using a fragment analyser such as the LabChip GXII RNA kit (or equivalent fragment analyser) following the manufacturer's instructions:
Expected result
The probes should have a peak at about 210-239 bp (with a tight distribution perhaps from 160-500 bp length).
Dilute the RNA probes to 200 ng/ul with RNAse-free water. Aliquot into 6 ul batches in 0.5 ml Lo-bind Safelock tubes and store at -80 deg C.
Label tubes.
| Tube | Qty | For ... | |
| 1.5 ml Lo-Bind Safelock tube | 1 | Mineral oil aliquot | |
| 0.2 ml Lo-Bind PCR tube | # capture reactions x 2 | Hybridisation | |
| 1.5 ml Lo-Bind Safelock tube | 2 | Step 14 and 15 master mixes |
Ensure all primer stocks are pepared in the correct buffer and that enough aliquots of the working concentrations are diluted out. Vortex and pulse centrifuge.
Thaw reagents. Flick all reagents to mix and pulse centrifuge where possible.
| Reagent | Stored at... | Thaw at... | |
| 20 X SSPE | 4 deg C | Room temperature | |
| 500 mM EDTA | 4 deg C | Room temperature | |
| 50 X Denardt's Solution | -20 deg C | Room temperature | |
| 10% SDS | 4 deg C | Room temperature | |
| Chickent Cot-1 (HyBloc) | -20 deg C | On ice | |
| FWD_blocking_primer | -20 deg C | On ice | |
| REV_blocking_primer | -20 deg C | On ice | |
| SUPERas-In RNAse inhibitor | -20 deg C | On ice | |
| Salmon Sperm DNA | -20 deg C | On ice |
Combine the following "BLOCKS" master mix in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | x ______ rxn | |
| Chicken Cot-1 (HyBloc) | 3.25 ul | 1 ug/ul | 0.77 ug/ul | 2.5 ul | ||
| Salmon Sperm DNA | 3.25 ul | 10 ug/ul | 0.77 ug/ul | 0.25 ul | ||
| FWD_blocking_primer | 3.25 ul | 200 uM | 15.4 uM | 0.25 ul | ||
| REV_blockg_primer | 3.25 ul | 200 uM | 15.4 uM | 0.25 ul |
"BLOCKS" master mix
Aliquot 3 ul of the BLOCKS master mix into 0.2 ml Lo-bind tubes, one for each capture reaction.
Combine the following "HYBS" master mix in a 1.5 ml Lo-bind Safelock tube. Vortex and pulse centrifuge.
| Reagent | V2 (reaction volume) | C1 (stock concentration) | C2 (concentration in reaction) | V1 (volume to add) | x ______ rxn | |
| SSPE | 20 ul | 20 X | 9 X | 9 ul | ||
| EDTA | 20 ul | 0.5 M | 0.0125 M | 0.5 ul | ||
| Denhardt's solution | 20 ul | 50 X | 8.75 X | 3.5 ul | ||
| SDS | 20 ul | 10 % | 0.25 % | 0.5 ul | ||
| SUPERase-In RNase inhibitor | 20 ul | 20 U/ul | 1 U/ul | 1 ul |
"HYBS" master mix
Aliquot 14.5 ul of the HYBS master mix into 0.2 ml Lo-bind tubes, one for each capture reaction.
Bring all the tubes to the post-PCR space.
Pre-program the thermal cycler:
95 deg C for 5 min
60 deg C for 5 min
60 deg C for hold
Note
Recommendations for the hybridisation temperature
Thaw a 6-ul aliquot of the probes from Step 10.13 on ice.
Add 5.5 ul of baits (ca. 500-1000 ng at 100-200 ng/ul) to each HYBS reaction. Flick to mix and pulse centrifuge. Place on ice.
Add 7 ul (100-500 ng) of shotgun library to the corresponding BLOCKS tube. Flick to mix and pulse centrifuge.
Note
Shotgun libraries can contain a single sample or a pool of samples...
Ideally the library should be double-indexed and purified (but not size-selected--a lot of the dimer should be washed away in the capture but it will be size-selected after the capture anyway).
The input amount of DNA is 100-500 ng in 7 ul (i.e., 14-72 ng/ul). The range is 1 ng input to up to 2 ug input DNA.
Transfer the BLOCKS tubes to a thermalcycler and start the program. Allow it to proceed through Step 1 (i.e., 95 deg C for 5 min).
When the thermalcycler reaches Step 2, transfer the HYBS tubes to the thermalcyler. Allow it to proceed through Step 2 (i.e., 60 deg C for 5 min).
When the thermalcycler reaches Step 3, transfer 18 ul from the HYBS tubes to the corresponding BLOCKS tubes. Pipette to mix. Discard the HYBS tubes.
Add 15 ul of mineral oil to the top of each reaction.
Allow the thermalcycler to proceed through Step 3 for 42 hr (i.e., 60 deg C for 42 hr).
Label tubes.
| Tube | Qty | For ... | |
| 50 ml Falcon tubes | 3 | Wash buffer 2, Wash buffer 2.2, Binding buffer | |
| 15 ml Falcon tubes | 3 | Aliquots for Wash buffer 2.2 and Binding buffer, and 10 mM Tris-HCl/0.05% Tween-20 | |
| 1.5 ml Lo-bind Safelock tube | 1 | Aliquot for MyOne C1 Streptavidin beads | |
| 1.5 ml Lo-bind Safelock tube | 3 x # hybridisation reactions | Enrichment |
Prepare all necessary buffers and UV decontaminate.
| Buffer | Reagent | Volume | C2 | |
| Wash buffer 2 | 20 X SSC | 100 ul | 0.1 X | |
| 10% SDS | 200 ul | 0.1 % | ||
| Ultrapure water | 19.7 ml | na | ||
| Wash buffer 2.2 | 10% SDS | 400 ul | 0.08% | |
| Wash buffer 2 | 10 ml | na | ||
| Ultrapure water | 39.6 ml | na | ||
| Binding buffer | 5 M NaCl | 10 ml | 1 M | |
| 1 M Tris-HCl | 500 ul | 10 mM | ||
| 0.5 M EDTA | 100 ul | 1 mM | ||
| Ultrapure water | 39.4 ml | na | ||
| 10 mM Tris-HCl, 0.05% Tween-20 | 1 M Tris-HCl | 500 ul | 10 mM | |
| 100% Tween-20 | 25 ul | 0.05% | ||
| Ultrapure water | 49.475 ml | na |
Aliquot reagents for to take to the post-PCR space.
| 700 ul * # reactions | Binding buffer | |
| 30 ul * # reactions | MyOne C1 Streptavidin beads | |
| 35 ul * # reactions | 10 mM Tris-HCl/0.05% Tween-20 | |
| 1600 ul * # reactions | Wash Buffer 2.2 |
Set a water bath or thermalshaker to 55 deg C. Warm Wash buffer 2.2 to 55 deg C for 45 min.
Set a heat block to 95 deg C.
Pellet the MyOne C1 Streptavidin beads for 2 min with the magnetic rack. Discard the supernatant.
Add 200 ul * # reactions of Binding buffer to the beads. Vortex and pulse centrifuge.
Pellet the beads with the magnetic rack and discard the supernatant.
Repeat Step 34-35 two more times for a total of 3 washes.
Resuspend the beads in 70 ul * # reactions of Binding buffer.
Aliquot 70 ul of beads into a 1.5 ml Lo-bind Eppendorf tube (1 per reaction).
Warm the bead aliquots to 55 deg C in the thermoshaker/water bath for 2 minutes.
Transfer the hybridised libraries at 60 deg C to the bead aliquots. Pipette to mix.
Incubate the hybridised libraries and beads in the thermoshaker/waterbath for 30 min at 55 deg C. Agitate every 5 minutes by flicking or gently continuously shake.
Pulse centrifuge. Pellet the beads with the magnetic rack. Discard the supernatant.
Add 500 ul heated Wash Buffer 2.2 to the beads. Vortex and pulse centrifuge.
Incubate 10 min at 55 deg C in the thermoshaker/water bath. Agitate every 2 min by flicking.
Pulse centrifuge. Pellet the beads with the magnetic rack. Discard the supernatant.
Repeat the wash steps above (Steps 43-45) two more times for a total of 3 washes.
Add 30 ul of 10 mM Tris-HCl/0.05% Tween-20 to the washed beads. Resuspend by pipetting.
Incubate at 95 deg C in a heat block for 5 min.
Pellet the beads with a magnetic rack and transfer the supernatant to a clean 1.5 ml tube.
At this point you can treat with RNAseA to remove any RNA or put RNAse A in the PCR but it is not necessary as the RNA will not amplify.
Dilute the captured libraries 1 in 10 in Ultrapure water (i.e., 1 ul in 9 ul Ultrapure water). Vortex and pulse centrifuge.
Label tubes.
Thaw reagents on ice.
Ensure that all primer stocks are prepared in the correct buffer and that enough aliquots of the working concentrations are diluted out. Vortex and pulse centrifuge.
| Oligo | Reagent | Volume | |
| 10 uM P5_primer | 100 uM P5 | 50 ul | |
| Ultrapure water | 450 ul | ||
| 10 uM P7_primer | 100 uM P7 | 50 ul | |
| Ultrapure water | 450 ul |
Make up the following master mix in a 1.5 ml Lo-bind tube.
| 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 | 25 ul | 10 uM | 0.4 uM | 1 ul | ||
| P7 | 25 ul | 10 uM | 0.4 uM | 1 ul |
Master mix for end-point PCR
Add 22.5 ul of master mix to each PCR tube following the schematic below. Pulse centrifuge the tubes.
| Capture001 Neat | ||
| Capture001 1in10 | ||
| Capture002 Neat | ||
| Capture002 1in10 | ||
| Capture003 Neat | ||
| Capture003 1in10 | ||
| PCR NTC | ||
| PCR NTC |
Make up the following master mix in a 1.5 ml Lo-bind tube.
| 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 | 25 ul | 10 uM | 0.4 uM | 1 ul | ||
| P7 | 25 ul | 10 uM | 0.4 uM | 1 ul |
Master mix for final PCR
Add 20 ul of master mix to each PCR tube following the schematic below. Pulse centrifuge the tubes.
| Capture001 Neat | Capture003 Neat | |
| Capture001 Neat | Capture003 Neat | |
| Capture001 Neat | Capture003 Neat | |
| Capture001 Neat | Capture003 Neat | |
| Capture002 Neat | ||
| Capture002 Neat | ||
| Capture002 Neat | ||
| Capture002 Neat |
Pulse centrifuge the 8-well qPCR strip tubes containing the master mix. Bring to the post-PCR space.
To the end-point PCR, add 2.5 ul of both neat and 1in10 captured library to the corresponding tubes. Add 2.5 ul nuclease free water to the remaining wells as PCR no-template controls. Vortex and pulse centrifuge.
Place tubes in the thermal cycler and run the following PCR 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
When the PCR is finished, determine the optimal number of cycles to give in the final PCR to ensure libraries are not over-amplified. i.e., stop the PCR during the linear phase of amplification.
Note
This is particularly important if you performed a capture that contained a pool of samples, and why if you DO capture a pool you ensure that both forward and reverse indexes are unique (not just the combination)--over-amplification can cause tag-jumping, so if the indexes are completely unique to a sample, even if tag jumping occurs they will be thrown out becuase the erroneous combination can be identified.
This step could be performed with alongside standards of known concentration to calculate the number of library molecules output from the capture library. This could be compared to to the number of library molecules input into the capture--if the capture worked, you would expect much fewer molecules OUT of the capture than went in.
The Neat and 1in10 dilution of libraries should show be approximately 3.33 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.
Agarose gel electrophoresis
Run 10 ul of PCR product from the first PCR on a 2% agarose gel electrophoresis.
Protocol
NAME
2% Agarose Gel Electrophoresis
CREATED BY
Alicia Grealy
To the second PCR add 5 ul of the neat library (as long as amplification was NOT inhibited in the first PCR; if amplification efficiency was poor, amplify a dilution and do more replicates). Perform enough replicates to amplify the entire captured library.
Perform Step 61 above, but stop the PCR during the linear phase of amplification as determined by the end-point PCR above.
Expected result
The number of cycles needed may be greater than was needed for the initial library amplification (before capture) but is usually between 15-20 cycles.
Combine replicates in a clean 1.5 mL Lo-bind Safelcok tube. Vortex and pulse centrifuge.
Purify
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:
Elute in 40 ul of Ultrapure water.
Run each enriched library 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:
Purify
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:
Elute in 25 ul of Ultrapure water.
Qubit
Measure the concentration of the neat library and these dilutions in duplicate on the Qubit following the manufacturer's instructions.
Dilute the libraries to 5 ng/ul in Ultrapure water in a total volume of 5-10 ul.
Fragment analyse
Use a LabChip GXII or equivalent fragment analyser (HiSense kit) to measure the molarity of the libraries between 160-500 bp.
Expected result
Libraries will be insert size + 136 bp, so the smallest fragments of interest will be ca. 166 bp (30 bp insert).
Pool enriched libraries in equimolar concentrations.
Dilute the libraries 1/2, 1/5, 1/10 in Ultrapure water (i.e., create a serial dilution in 10 ul volume).
Qubit
Measure the concentration of the neat library and these dilutions in duplicate on the Qubit following the manufacturer's instructions.
Fragment analyse
Measure the molarity of the neat library and dilutions on a LabChip GXII Hisense kit (or equivalent fragment analyser) following the manufacturer's instructions:
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.
Dilute the library to between 2-4 nM in Ultrapure water.
Note
Note that if your libraries were built using the single-stranded protocol (Gansauge and Meyer 2013; Gansauge et al. 2017), you will need CL72_custom_sequencing_primer to sequence. This can be spiked into well 12 (but select 'no custom primer' in the run set up) or into well 18 (select 'custom primer' in the run set up). Spiking the custom primers into the run is preferable so that the remaining Illumina primers are present and can sequence PhiX.
You do not need custom i5_indexing_primer to sequence off a MiSeq or NovaSeq because these instruments prime off P5. You do not need a custom i7 indexing primer because it uses primers already included in the kit. Note that for the NextSeq you will need custom i5_indexing_primer in addition to the CL72_custom_sequencing_primer.
Follow the manufacturer's instructions to perform the sequencnig run on your Illumina platform of choice.
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.
Alternatively, concentrate the libraries using a SpeedyVac system, following the manufacturer's instructions.
Size select and purify
Size select and purify
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:
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:
Elute in 25 ul of Ultrapure water.
Quantitate the final library
Quantitate the final library
Dilute the libraries 1/2, 1/5, 1/10 in Ultrapure water (i.e., create a serial dilution in 10 ul volume).
Measure the concentration of the neat library and these dilutions in duplicate on the Qubit following the manufacturer's instructions.
Measure the molarity of the neat library and dilutions on a LabChip GXII Hisense kit (or equivalent fragment analyser) following the manufacturer's instructions:
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
Sequencing
1h
Dilute the library to between 2-4 nM in Ultrapure water.
Follow the manufacturer's instructions to perform the sequencnig run on your platform of choice.