Ovation RRBS Methyl-seq library preparation protocol

Prepare a list of samples and their concentrations, calculate the required volumes for 100ng DNA and nuclease-free water for a final reaction volume of 8.5µL.

Calculate the volume of a methylation conversion control to spike in each sample. This is usually unmethylated Lambda virus DNA, added in 1:1000 – 1:5000 ratio to each DNA sample – i.e. up to 0.1ng unmethylated DNA control per sample. The starting unmethylated DNA stock will be highly concentrated and a dilution will be required.

Prepare the unmethylated DNA control – add the necessary total volume to the total volume of water (from the calculations, sum up the unmethylated control DNA and water volumes for all samples), and pre-mix well.

Add water (+ unmethylated control) into 0.2 mL PCR strip tubes (if doing 24 samples) or a plate - usually starting with the highest amount of water going down the sample list, column after column. Cross-check for each with the order on the plate schematic, re sample ID, required volume and correct well position.

Add DNA samples in the same sample order, column by column, starting from low to high volume. Cross-check for each with the order on the plate schematic, re sample ID, required DNA volume and correct well position. It is a good practice to cover the completed plate columns as going forward. Keep DNA samples On ice.

Thaw MspI Buffer Mix at Room temperature. Mix MspI Buffer Mix by vortexing, spin and place On ice.

Spin down the MspI Enzyme Mix and place On ice.

Prepare a master mix by combining MspI Buffer Mix and MspI Enzyme Mix in a 0.2ml PCR tube within an 8-tube PCR strip, according to the volumes shown in Table 1. Mix by pipetting, spin down briefly and immediately place On ice.

Table 1. MspI Master Mix (advisable extra volume included for high sample number preps).

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME (55)	96X RXN VOLUME (110)	STORAGE
MspI BUFFER MIX (BLUE)	1.0 μL	54 μL	110 μL	–20°C
MspI ENZYME MIX (BLUE)	0.5 μL	27 μL	55 μL	–20°C

Distribute equal amounts of master mix to the 8 tubes in the PCR strip for multichannel pipetting.

Add 1.5µL of MspI Master Mix to each sample tube for a total of 10µL (with a reliable 0.5-10 μL multichannel and low retention tips). Mix by pipetting (increase pipette volume for the mixing), spin down and place On ice.

Place the tubes in a thermal cycler programmed to run Program 1 (MspI Digestion):

• 37°C – 1h 0m 0s,
• hold at 4°C.

In the meantime take out to thaw at RT the water and buffers for adapter ligation – see Adaptor Ligation .

Remove the tubes from the thermal cycler, spin to collect condensation and place On ice.

Thaw Ligation Buffer Mix L1 at Room temperature. Mix well by vortexing, spin and place On ice.

Thaw the Ligation Adaptor Plate On ice, spin down, and return to ice.

Spin down the Ligation Enzyme Mix L3 and place On ice or in a cooling rack.

Pierce the Ligation Adaptor Plate seal with the tips and aspirate 3µL (the entire amount) of the Ligation Adaptor Mix L2 (with a 0.5-10ul multichannel and low retention tips). Add one adaptor barcode per sample, noting the adaptor plate positions per each sample. Mix thoroughly by pipetting and keep On ice.

Just prior to use, prepare a master mix by combining D1, L1 and L3, according to the volumes shown in Table 2. Mix by pipetting slowly, without introducing bubbles, spin and place On ice. Use the master mix immediately.

Table 2. Ligation Master Mix (advisable extra volume included for high sample number preps)

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME (55)	96X RXN VOLUME (105)	STORAGE
NUCLEASE-FREE WATER (GREEN: D1)	2.0 μL	110 μL	210 μL	-
LIGATION BUFFER MIX (YELLOW: L1)	4.0 μL	220 μL	420 μL	–20°C
LIGATION ENZYME MIX (YELLOW: L3)	1.0 μL	55 μL	105 μL	–20°C

Add 7µL Ligation Master Mix to each reaction tube for a total of 20µL (with a multi-dispenser or a reliable 0.5-10μl/2-20μl multichannel and low retention tips). Mix thoroughly by pipetting slowly and gently, spin down and place on ice. Proceed immediately with the incubation.

Place the tubes in a thermal cycler programmed to run Program 2 (Ligation):

• 25°C – 0h 30m 0s,
• 70°C – 0h 10m 0s,
• hold at 4°C.

Remove the tubes from the thermal cycler, spin to collect condensation and place On ice.

Remove the Magnetic Bead Solution and Binding Buffer 1 from 4°C and place at Room temperature for use in the next step.

Thaw Final Repair Buffer Mix (FR1) at Room temperature. Mix by vortexing, spin down and place On ice.

Spin down Final Repair Buffer Enzyme (FR2) and place On ice.

Prepare a master mix by combining FR1, FR2 and Nuclease-free Water (D1) according to the volumes shown in Table 3.

Table. 3 Final Repair Master Mix  (advisable extra volume included for high sample number preps)

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME (54)	96X RXN VOLUME (100)	STORAGE
FINAL REPAIR BUFFER MIX (PURPLE: FR1 ver 4)	6 μL	324 μL	600 μL	–20 °C
FINAL REPAIR ENZYME MIX (PURPLE: FR2)	0.5 μL	27 μL	50 μL	–20 °C
NUCLEASE-FREE WATER (GREEN: D1)	13.5 μL	729 μL	1350 μL	–

Add 20µL of the Final Repair Master Mix to each sample for a total of 40µL (use a reliable multichannel pipette and low retention tips). Mix by pipetting each time, spin down and place On ice.

Place the tubes in a thermal cycler pre-heated to 60°C (if possible) and programmed to run Program 3 (Final Repair):

• 60°C – 0h 10m 0s,
• 70°C – 0h 10m 0s,
• hold at 4°C.

Remove the tubes from the thermal cycler, spin to collect condensation and place On ice.

Remove Magnetic Bead Solution and Binding Buffer 1 from storage and place on bench top. Ensure they have reached Room temperature before use.

Once warmed to room temperature, mix Binding Buffer 1 by inversion until homogenized.

Vortex Magnetic Bead Solution until homogenized.

Prepare a master mix of Magnetic Bead Binding Solution 1 (MBBS1) as directed in Table 4. Binding

Buffer 1 is supplied in5.4mL bottles suitable for a 48 sample mix, use two bottles for a full plate.

Table. 4 Magnetic Bead Binding Solution 1 Master Mix.

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME (54)	96X RXN VOLUME (108)	STORAGE
BINDING BUFFER 1	100 μL	5.4 mL	2 x 5.4 mL	4 °C
MAGNETIC BEAD SOLUTION	2.0 μL	108 μL	2 x 108 μL	4 °C

Vortex MBBS1 master mix thoroughly to ensure the beads are homogenized in solution.

At Room temperature, add 10µL of Ultra Pure water to each sample for a total of 50µL (with a reliable 0.5-10 μl/2-20 μl multichannel and low retention tips or an Eppendorf dispenser, without touching the wells, 1.0 ml tip).

Add 100µL of MBBS1 master mix to each 0.2 mL tube/well containing 50µL sample for a total of 150µL (with a dispenser, 2-10 ml tip). Mix by pipetting with a multichannel pipette set at 130 μL and centrifuge briefly.

Incubate at Room temperature for 0h 20m 0s (can be longer if a break is needed).

Prepare a fresh stock of 80% ethanol, using the Ultra Pure water provided with the kit. Mix by inversion and place at Room temperature.

Transfer tubes/plate to a magnetic separation plate and incubate at Room temperature for 0h 10m 0s or until the solution of beads is completely clear.

Keeping the tubes on the magnet, carefully remove the supernatant and discard it – with a manual multichannel pipette set at 140µL.

With the tubes/plate still on the magnet, carefully add 200µL of 80% ethanol wash to the wells without disturbing the bead pellet (with a dispensing multi-channel set at 1200µL, 6x strips/half a plate at a time).

Remove and discard the 200µL 80% ethanol wash, avoiding aspiration of the bead pellet (with a manual 20-200 μL multichannel, regular tips). Complete this step as quickly as possible, ideally within 2 minutes.

Repeat Steps 40 and 41 to perform 2 x 200µL 80% ethanol washes in total. Remove as much of the final wash as possible – first with a 20-200 μL tip, and a second time with a 10/20 μL

tip if necessary.

Air dry the bead pellets for 5-10 minutes at Room temperature.

Remove the tubes from the magnet.

Add 11µL of Elution Buffer directly onto the bead pellet.

Mix thoroughly with a 0.5-10μl multichannel to ensure all beads are resuspended. Don’t centrifuge if there are drops on the walls!

Incubate at Room temperature for at least 0h 5m 0s to elute the TrueMethyl converted DNA from the beads.

Seal the plate and centrifuge briefly to collect all drops at the bottom of the tubes.

Transfer tubes/plate to a magnetic separation plate and incubate at Room temperature for 5-10 minutes or until the solution of beads is completely clear.

Removing the seal one column/strip at a time, carefully aspirate 10µL of the eluate, ensuring as few beads as possible are carried over, and transfer to a new microcentrifuge plate/tubes (use a 0.5-10uL multichannel with low retention tips). Keep at Room temperature if continuing with section Bisulfite Conversion . Can store in fridge or freezer if an 0h 5m 0s break is needed.

Set a heat block or heated orbital incubator to 70°C.

Remove Bisulfite Diluent and Bisulfite Reagent aliquots from storage and place on bench top. Remove 1 aliquot of Bisulfite Reagent for every 25 reactions to be processed and spin quickly to remove any powder from cap (i.e. a full 96-well plate uses 4 Bisulfite Reagent vials).

Prepare Bisulfite Reagent Solution by adding 700µL of Bisulfite Diluent to each aliquot of Bisulfite Reagent.

Seal the lid of each aliquot with Bisulfite Reagent Solution tightly.

Incubate the aliquots of Bisulfite Reagent Solution for minimum 0h 30m 0s at 70°C and vortex regularly until the Bisulfite Reagent Solution is completely (or nearly completely) dissolved. This step can take up to 3h 0m 0s for the reagent to dissolve.

Spin down Bisulfite Reagent Solution briefly and place at Room temperature.

Ensure purified DNA samples from previous step are at Room temperature before proceeding.

Prepare Bisulfite Conversion Reaction mix by adding 30µL of Bisulfite Reagent Solution to each 10µL of DNA for a total of 40µL (with multi-dispenser, 1 mL tips).

Mix by pipetting with a 20-200 μl multichannel, spin down and place at Room temperature.

Place the tubes in a pre-warmed thermal cycler programmed to run Program 5 (Bisulfite Conversion):

• 99°C – 0h 10m 0s,
• 65°C – 1h 45m 0s,
• hold at 20°C.
  Note

  Optional stopping point : You may hold samples at Room temperature (20°C) for up to 16h 0m 0s. Do not store below 20°C.

Once the bisulfite conversion is complete, centrifuge samples briefly to collect the solution at bottom of the tubes.

Continue to Bisulfite-Converted DNA Desulfonation and Purification .

Remove Desulfonation Buffer, Binding Buffer 2, Magnetic Bead Solution and Elution Buffer from storage and place at Room temperature for a minimum of 0h 30m 0s before use.

Prepare a fresh stock of 80% Ethanol. Mix by vortexing or inversion.

Mix Binding Buffer 2 by inversion until homogenized.

Vortex Magnetic Bead Solution until homogenized.

Prepare a master mix of Magnetic Bead Binding Solution 2 (MBBS2) as directed in Table 5. Each kit BB2 bottle is filled with exactly 9mL solution ready to use for a 48 sample prep; use two bottles for a full plate.

Table. 5 Magnetic Bead Binding Solution 2 Master Mix.

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME	96X RXN VOLUME	STORAGE
BINDING BUFFER 2	160 μL	9 mL	2 x 9 mL	4 °C
MAGNETIC BEAD SOLUTION	1.92 μL	108 μL	2 x 108 μL	4 °C

Vortex MBBS2 thoroughly to ensure the solution is homogenous before aliquoting.

Carefully add 160µL of MBBS2 to each tube containing 40µL of bisulfite converted sample for a total of 200µL (with multi-dispenser, 5 mL tip). Mix thoroughly by pipetting slowly and gently, spin down and place at Room temperature.

Incubate at Room temperature for 0h 30m 0s to 1h 0m 0s, vortexing if necessary, or alternating plate position by rotation upwards-downwards after sealing VERY WELL.

Prepare desulfonation buffer by adding 7mL 100% ethanol to 3mL of Desulfonation Buffer for up to 48 samples, or 14mL EtOH + 6mL Desulfonation buffer for up to 96 samples, respectively.

Centrifuge briefly to collect the solution at the bottom of the tubes/wells. To bring down the beads from the liquid remaining above the magnet, centrifuge at 2200 rcf / 4700rpm.

Place the tubes/plate onto the magnet and incubate at Room temperature for 10-15 minutes to completely clear the solution of beads.

Carefully remove the supernatant and discard it, without discarding any beads.

Add 200µL of 80% Ethanol to each sample tube/well (with a dispensing multichannel up to 1200µL), 6 columns at a time).

Carefully remove the 80% Ethanol wash and discard it. Remove as much of the wash as possible.

Add 200µL of Desulfonation Buffer with added Ethanol onto the bead pellet. Incubate at Room temperature for 0h 5m 0s.

Carefully remove 200µL of the Desulfonation Buffer and discard it. Remove as much of the Desulfonation Buffer as possible without disturbing the bead pellet.

With the tubes/plate still on the magnet, add 200µL of 80% Ethanol to each sample tube without disturbing the bead pellet and incubate for 0h 0m 30s (with a dispensing multichannel set at 1200 μL, 6x strips/half a plate at a time).

Remove the 200µL 80% Ethanol wash and discard it, avoiding aspiration of the bead pellet (with a manual 20-200 μL multichannel, regular tips). This step must be completed as quickly as possible (ideally within 0h 2m 0s)

Repeat steps 79–80 to perform 2 x 200µL 80% Ethanol washes in total. Remove as much of the final wash as possible – first with a 20-200 μL tip, and a second time with a 10/20 μL tip, if necessary.

Air-dry the the beads on the magnet for 5-10 minutes. Inspect each tube carefully to ensure that all of the Ethanol has evaporated – the pellet should be non-shiny (matt look) when it is ready. Using a pipette with 10 μl tips can help with aspirating Ethanol drops while still on the magnet.

Remove the tubes/plate from the magnet.

Quickly add Elution Buffer with a multi-dispensing pipette aiming directly onto the bead pellet:

For Library Amplification Optimzation with qPCR (recommended), resuspend the beads in 25µL of Elution Buffer.

If qPCR optimization is not required, resuspend the beads in 21µL of Elution Buffer.

Mix and resuspend completely the beads in the Elution Buffer with a multichannel pipette pipette – make sure all beads are removed from the well walls and brought in solution.

Incubate at Room temperature for 0h 5m 0s to elute the bisulfite converted DNA from the beads.

Seal the plate and centrifuge briefly to collect the samples at the bottom of the tubes/wells.

Place the tubes/plate onto the magnet and incubate at Room temperature for 5-10 minutes to completely clear the solution of beads.

Opening the seal one column/strip at a time, carefully transfer eluate into a fresh 0.2 mL PCR plate:

For Library Amplification Optimzation with qPCR (recommended first time only), transfer 24µL of eluate.

If qPCR optimization is not required, transfer 20µL of eluate.

Prepare a master mix by combining P2, P3 and 20x EvaGreen in an appropriately sized capped tube according to the volumes shown in Table 6. Add P3 at the last moment and mix well by pipetting, taking care to avoid bubbles. Spin down and place On ice.

Table. 6 Library Amplification qPCR Master Mix (with extra volumes for high sample number).

A	B	C	D	E
REAGENT	1X RXN VOLUME	4X RXN VOLUME	96X RXN VOLUME	STORAGE
AMPLIFICATION PRIMER MIX (RED: P2 ver 8)	1.0 μL	4.2 μL	100 μL	–20 °C
AMPLIFICATION ENZYME MIX (RED: P3 ver 3)	4.5 μL	18.9 μL	450 μL	–20 °C
20X EvaGreen	0.5 μL	2.1 μL	50 μL	–20 °C

Aliquot 6µL of master mix per sample into an appropriate optically clear PCR plate. Spin down and place On ice.

On ice, add 4µL of sample to each 6µL of Library Amplification qPCR Master Mix for a total of 10µL per reaction. Keep the remaining 20µL of sample On ice.

Mix well by pipetting, spin down and place On ice.

Perform qPCR with the following cycling conditions:

• 95°C – 0h 2m 0s,
• 35 x (95°C– 0h 0m 15s, 60°C – 0h 1m 0s, 72°C – 0h 0m 30s).

Examine the log fluorescence vs. cycle number plot from the qPCR system to determine the appropriate number of library amplification cycles. Select a cycle number within the middle to late exponential phase of the amplification plot. Examples are provided in Figure 8 of the original Tecan kit manual.

Remove Agencourt Beads from 4°C and DR1 from -20°C and place at Room temperature for use in the next step.

Spin down Amplification Enzyme Mix (P3) and place On ice.

Thaw Amplification Primer Mix (P2) at Room temperature. Mix by vortexing, spin down and place On ice.

Prepare a master mix by combining P2 and P3 according to the volumes shown in Table 7. Mix well by pipetting, taking care to avoid bubbles, spin down and place On ice.

Table. 7 Library Amplification Master Mix (with extra volumes for high sample number) .

A	B	C	D	E
REAGENT	1X RXN VOLUME	48X RXN VOLUME (50)	96X RXN VOLUME (100)	STORAGE
AMPLIFICATION PRIMER MIX (RED: P2 ver 8)	5.0 μL	275 μL	500 μL	–20 °C
AMPLIFICATION ENZYME MIX (RED: P2 ver 3)	25.0 μL	1.375 μL	2.500 μL	–20 °C

On ice, add 30µL of Amplification Master Mix to each sample for a total of 50µL.

Place tubes in a pre-warmed thermal cycler programmed to run Program 6 (Library Amplification):

• 95°C – 0h 2m 0s,
• N(95°C – 0h 0m 15s,
• 60°C – 0h 1m 0s,
• 72°C – 0h 0m 30s),
• 72°C – 0h 5m 0s,
• hold at 10°C.

Remove the tubes from the thermal cycler, spin to collect condensation and place On ice.

Ensure the Agencourt beads and DR1 Resuspension Buffer have reached Room temperature before proceeding.

Resuspend the beads by inverting and tapping the tube. Ensure the beads are fully resuspended before adding to samples. After resuspending, do not spin the beads.

Add 50µL (1 volume) of the bead suspension to each reaction. Mix thoroughly by pipetting 10 times.

Incubate at Room temperature for 0h 10m 0s.

Prepare fresh 80% EtOH solution in a 50 mL falcon tube – 40mL for a full plate, 20mL for a half plate.

Transfer the tubes to the magnet and let stand 0h 5m 0s to completely clear the solution of beads.

If doing your own QC, prepare a plate for Tapestation aliquots and take out Tapestation kit reagents to equilibrate to Room temperature (see Library Quality Control step).

Carefully remove 90µL of the binding buffer and discard it. Leaving some of the volume behind minimizes bead loss at this step.

With samples still on the magnet, add 200µL of of 80% ethanol to each sample tube and incubate for 0h 0m 30s (with a dispensing multichannel set at 1200 μL, 6x strips/half a plate at a time).

Remove the 200 μL of 80% Ethanol wash and discard it (with a manual 20-200 μL multichannel, regular tips). This step must be completed as quickly as possible (ideally within 2 minutes).

Repeat steps 111 and 112 for a total of two washes. Remove all remaining traces of ethanol after the second wash.

Air dry the beads on the magnet for 5-10 minutes. Inspect each tube carefully to ensure that all of the ethanol has evaporated. It is critical that all residual ethanol be removed prior to continuing, but it is also critical to not let beads over-dry as this reduces yield (they have over-dried when they crack).

Remove the tubes from the magnet and add 20µL DR1 Resuspension buffer to the dried beads (with dispenser aiming at the beads).

Mix thoroughly with a multichannel pipette, set at 10µL, to ensure all beads are resuspended – if the beads have cracked this may require mixing for a few minutes. Take any beads remaining on the sides and make sure all beads are covered in solution.

Seal the plate/close the tubes and centrifuge briefly to collect sample at bottom of the tubes/wells.

Transfer the tubes to the magnet and let the samples stand for 0h 5m 0s or until the solution is completely clear of beads.

Carefully remove the seal one column/strip at a time, and aspirate 19µL of the eluate, ensuring as few beads as possible are carried over, and transfer to a fresh plate / set of PCR tubes. Keep the eluate on ice.

Once the elution is complete, proceed with preparing library aliquots for a Tapestation QC run in the next section Library Quality Control (QC) . If QC is not planned soon, seal the library plate well and store at -20°C .

Aliquot 1µL of nuclease-free water in the wells of a 96-well plate, matching the number and positions of eluted libraries (this is for a 1 in 2 dilution, respectively, higher dilutions can also be used, depending on QC equipment sensitivity).

From the elution plate from Step 120, take 1µL of purified library with a multichannel pipette, transfer to the QC plate with water and mix.

The plate with purified libraries can now be sealed and stored long term at -20°C.

If performing a Tapestation run, continue with the High Sensitivity DNA 1000 Tapestation kit protocol (procedure not described here). If taking the DNA to be measured elsewhere, seal the QC plate and take to the facility. The QC plate can be stored at -20°C, although evaporation can occur during long term storage.