DEACYLATED tRNA Library Preparation for RNA004 Sequencing (FINAL)
Introduction
This protocol will take you through the optimized deacylated tRNA library preparation for nanopore sequencing using the RNA004 chemistry. Before you begin this protocol, you must extract RNA from your species of interest and acquire 3-10 µg of total RNA.
GOOD RNA PRACTICES:
You need to be working in an RNase-free environment. Unless stated otherwise, RNA and reagents should be kept on ice at all times. To store overnight, RNA is always frozen at -80ºC.
INSTRUCTIONS ON ACCESS:
This protocol will be made available via a PDF and a Benchling link. Anyone can open the shared link and view the protocol even when they are signed out of a Benchling account (or don't have one). This viewer will allow you to edit the tables only in a way that will not impact the master copy. You will not be able to edit the text on this viewer. The viewer also has a step-by-step feature which allows one to check off completed steps and start timers. This is probably the best electronic way to access the protocol if you do not use Benchling.
If you do use Benchling and want to add this protocol to your personal protocols, you can also do that. Just open the shared link while signed into your Benchling account. You will see a clock symbol underneath the dark blue share symbol in the top right margin of the protocol. You will see verion history here. Select the most recent version and click clone from version in the bottom right of this popup. You can then select one of your folders to clone the protocol into. Once you've done that, you now have your own copy that you can edit and add to entries as you please.
USEFUL RESOURCE: I like this
online calculator for converting between moles and grams for nucleic acids.
Materials
3-10 µg of isolated total RNA, quantified on Nanodrop
DEPC treated, nuclease free water (Ambion)
100% and 80% ethanol
Zymo RNA Clean and Concentrator-5 kit (Zymo Research, R1016)
10 µM pre-annealed tRNA splint adapter (48 nt combined)
PEG 8000 (NEB B1004S)
T4 RNA ligase 2 and its buffer (NEB)
5X Quick ligation buffer (NEB B6058S)
T4 DNA ligase (NEB, M0202M, 2,000,000 U/mL)
RNase inhibitor (Enzymatics or equivalent)
BioDynami tRNA beads (40054S)
Unlike single stranded RNA oligos of equivalent size, tRNAs do not readily bind SPRI beads. The Hesselberth lab has tested these beads for purifying ligated tRNAs, and confirmed their capacity to purify tRNAs and tRNA + adapter ligation products over a 75 - 177 nt range.
Ampure XP beads (Fisher Scientific A63881)
ONT sequencing kit (contains RTA adapter, though we can also generate in house RTA mixes)
ONT flow cell priming kit
HS DNA 1000 Qubit reagents
Procedure
Your sample needs to be between 3 and 10 µg total. You need this amount in a 25 µL volume of water (or equivalent).
Add 25 µL of 200 mM Tris pH 9.0 to your tRNAs. The final concentration of Tris pH will be 100 mM.
Incubate for 30 minutes at 30ºC.
Use the RNA clean and concentrate kit 5 according to the manufacturer's instructions. They are copied below. Only the green step has been adjusted silghtly from the manufacturer's instructions.
Mix 50 µL of RNA Binding Buffer with equivolume 100% ethanol.
Add the 100 µL of the above solution to your deacylation reaction.
if your deacylation reaction is not 50 µL, make sure you are adding 2 volumes of the 100% EtOH + RNA binding buffer to your deacylated tRNAs.
Transfer to a Zymo-IICR column COLLECTING IN AN EPPENDORF and centrifuge. Your small RNA sample will be in the flow through, do not discard it!
Add a 1.3X volume of 100% ethanol to the flow through. If you started with 150 µL tRNAs + EtOH + RNA binding buffer, add 195 µL 100% EtOH.
The manufacture's instructions say a 1X volume instead of a 1.3X volume.
Transfer to a new Zymo-IC column and centrifuge for 30 seconds at 10000 rpm.
Add 400 µL RNA prep buffer to the column and centrifuge for 30 seconds at 10000 rpm. Discard flow-through.
Add 700 µL of RNA wash buffer and centrifuge for 30 seconds at 10000 rpm. Discard flow-through.
Add another 400 µL RNA wash buffer and centrifugefor 30 seconds at 10000 rpm. Discard flow-through.
Spin once more for 5+ minutes at 10000 rpm to dry column.
Transfer to a fresh collection tube, add 11.5 µL nuclease free water directly to the column matrix. Centrifuge for 30 seconds at 10000 rpm to elute.
Nanodrop the sample using 1 µL.
Samples with low 260/230 (<1.75) should always undergo additional cleanup before library prep. If your 260/230 is especially low, you may have EtOH in your elution. You can potentially fix this by doing another cleanup with the RNA clean and concentrate kit 5 following the instructions for "Total RNA clenaup".
Ligate deacylated tRNAs to the splint adaptors. The optimal molar ratio for this ligation is 1 pmol tRNA: 1 pmol splint adapters.
Dilute your sample such that you have 10 µL water containing 300 ng deacylated tRNAs (30 ng/µL). This is an important step to be precise on.
The ligation setup below is for a 10 µL sample containing 300 ng tRNAs (30 ng/µL) input ONLY!
If your sample deviates from this amount, you need to use the table in Appendix 2 to calculate volumes for the ligation in order to maintain the optimal ratio of tRNAs to splint adapters.
Set up the following 20 µL reaction. Add enzymes to master mix last.
The master mix will be 120% of the volume you need to account for pipetting error.
tRNA dilutions for splint adapter ligation
Splint adaptor ligation set up
Incubate for 30 minutes at RT.
This used to be 1 hour, but we determined 30 minutes is just as efficient.
Equlibrate BioDynami tRNA beads to RT. Below I have copied the BioDynami protocol with some minor optimizations that we made to save time.
Add 36 µL (1.8X volume) of WELL-VORTEXED beads to the ligation reaction.
Mix by pipetting and incubate at RT for 4 minutes.
Incubate for 2 more minutes or until the beads are all gathered on the magnet.
Wash without disturbing the beads with 200 µL 80% EtOH.
Incubate for 1 minute and remove as much of the EtOH as possible. If you remove what you can, spin down in a benchtop centrifuge, resettle the beads on the magnet, you'll be able to remove more EtOH.
Dry with cap open until no EtOH is visible. This usually take 5-10 minutes if you've done a good job removing what you can.
Resuspend in 13 µL water UNLESS YOU USED APPENDIX 2 FOR THE SPLINT ADAPTER LIGATION. If you used Appendix 2, elute in 14 µL.
Incubate for 1 minute or until beads are settled on the magnet and remove the elution to a new tube.
You can expect to get a yield of 5-6 pmol ligated tRNA (27-33 ng/µL in 9 µL or 250-300 total ng) for a 300 ng input into the splint adapter ligation. This is quite consistent so we don't quantify again after splint adapter ligation if you used 300 ng tRNAs as input.
ONLY IF YOU USED APPENDIX 2 FOR SPLINT ADAPTER LIGRATION, nanodrop 1 µL.
Ligate the splint-adapter-ligated tRNAs to the RTA adapter. The optimal molar ratio is 3 pmol tRNAs with splint adapters: 1 pmol RTA adapter.
If you did NOT use the prescribed 300 ng tRNAs as input for the splint adapter ligation, you MUST use the table in Appendix 3 to recalculate your volumes for this ligation. The volumes given here are ONLY for those that used 300 ng tRNAs in the splint adaptor ligation.
Choose the table below for the T4 DNA ligase you have. We have been successful with both reactions below.
Set up the following 20 µL ligation. Add enzyme last.
Note: The RTA adapter was determined to be 1.4 µM by our lab.
RTA Adaptor Ligation Set up
Incubate for 30 minutes at RT.
If this is your lab's first time doing this protocol, you may want to look at your samples by Tapestation after the RTA ligation. If you are planning on doing this, take out HS DNA 1000 reagents to equilibrate to RT. If you do not want/need to Tapestation take out Quant-iT Qubit dsDNA HS reagents out to equilibrate to RT.
Quantification can be done another day if you are tight on time, so obvisouly only get out reagents if you plan on quantifying same day.
Equlibrate BioDynami tRNA beads to RT. Below I have copied the BioDynami protocol with some minor optimizations that we made to save time.
This process is the same as the cleanup after splint adapter ligation except for the volumes in yellow.
Add 27 µL (1.35X volume) of WELL-VORTEXED beads to the ligation reaction.
Mix by pipetting and incubate at RT for 4 minutes.
Incubate for 2 more minutes or until the beads are all gathered on the magnet.
Wash without disturbing the beads with 200 µL 80% EtOH.
Incubate for 1 minute and remove as much of the EtOH as possible. If you remove what you can, spin down in a benchtop centrifuge, resettle the beads on the magnet, you'll be able to remove more EtOH.
Dry with cap open until no EtOH is visible. This usually take 5-10 minutes if you've done a good job removing what you can.
Resuspend in 25 µL water. This is a different volume than last time.
Incubate for 1 minute or until beads are settled on the magnet and remove the elution to a new tube.
Nanodrop to roughly quantify with 1 µL.
You can expect 1 pmol (~2.4 ng/µL in 25 µL, 60 ng total) of ligated material.
You have two options to quantify more accurately: Tapestation or Qubit. Here's how to choose which method is best:
Qubit if you got a reading between 5 and 20 ng/µL on the nanodrop. Your lab has done this workup before and is familiar with it.
Tapestation if this is your first time through this protocol or your nanodrop reading falls outside of the above range.
Make sure the reagents have eqiulibrated to RT. This includes dsDNA HS buffer and dsDNA HS standards (1 and 2).
Get out the number of qubit tubes you need (number of samples + 2).
Set up the following master mix.
Add 10 µL of each standard to a Qubit tube.
Add 1 µL of each of your samples (should be roughly 10 ng, but anywhere from 0.2-100 ng is acceptable).
Add 190 µL of working solution to each standard and 199 µL to each sample.
Vortex all tubes and let sit for 2 minutes.
Read the concentration of the tubes and convert from ng/µL to pg/µL by multiplying by 1000.
Make sure reagents are equilibrated to RT.
Use 1 µL of each of your samples to make a working solution that is 0.01 - 1 ng/µL. Usually a 1:10 dilution in water will be good.
Run a HS DNA 1000 tape. The instructions are copied below.
Put the tape in the Tapestation. Always use up the remaining slots on an opened tape before starting a new tape.
Add 2 µL High Sensitivity D1000 Sample Buffer to 2 µL of your working solution in Tapestation tubes.
Vortex at 2000 rpm for 1 minute.
Place tubes in the Tapestation loading tray. Make sure enough tips are in the Tapestation. Run the program.
You should expect to see peaks at ~150 nts and ~300 nts. The expected size of the RTA-ligated products is ~177 nts, so you should use the estimated amount in the ~150 nt peak as your quantification. Don't worry about the 300 nt peak. We never see it come out in sequencing, it may just be an artifact.
Convert the concentration of your RTA-ligated tRNAs to pg/µL. The table will take into account any dilutions you made so don't do that yourself. Instead, input the quantification exactly as it was given to you by Qubit or the Tapestation.
Enter this concentration in the table below. Also add the dilution factor used when quantifying. Since we used 1 µL of product either for Qubit or to dilute for Tapestation, the dilution factor will be equal to the total volume of the working solution used for quantification. If you didn't deviate from the instructions here, your value is 1 if you Qubited and 10 if you ran a tape.
Add the remaining volume of your sample.
You will need between 50 and 400 fmol of product for the RLA ligation. You must be in this range. If you are not, we can't say how your sequencing will look if you proceed. Starting with 300 ng of input at the splint adapter step should ensure that you have enough product. We typically go forward with 100 fmol in 23 µL total. The table below will calculate your total yield as well as the volume of sample and water you must add to a new tube to make a 23 µL solution with 100 fmol.
Calculation of RTA-ligated yeild
Make 100 fmol 23 µL solutions. If you aren't going to sequence TODAY, stop here and freeze at -80ºC until the day you will sequence.
You should not continue this protocol unless you have time to finish it and sequence TODAY.
Set up the following 40 µL ligation of the helicase-loaded adapter. Keep the table with your T4 DNA ligase of choice.
The RLA adapter is the most expensive reagent in your sequencing kit, so don't waste it!
RLA Adapter Ligation Setup
On ice, start thawing tubes of Sequencing Buffer (SB), Library Solution (LIS), RNA Flush Tether (RFT) and Flow Cell Flush (FCF). Also start thawing flow cell wash reagents: wash mix (WMX), wash diluent (DIL), and storage buffer (S). FCF, DIL, and S are often in larger sizes and may take longer to thaw, so be aware of that.
Incubate for 30 minutes at RT.
Start equilibrating flow cell to RT (20 minutes equilibration needed). Check for condensation on underside and gold connector pins and wipe any off. This is more likely to be necessary in humid climates.
Equlibrate Ampure XP beads to RT.
Ampure XP beads struggle to bind tRNA efficiently, but at this point, the adapters make up enough of the total molecule that they can now bind the ligated products.
Add 72 µL (1.8x volume) Ampure XP beads.
We are using a 1.8X volume of beads instead of ONT's recommened 0.4X volume because tRNAs are smaller than most of the RNAs you would sequence. We need this higher bead volume to help us selectively bind tRNAs and not the 90 nt free RLA adapter.
Wash twice with 150 µL WSB (ONT) resuspending beads by flicking the tube and then reseating on the magnet to settle. Remove the wash buffer each time after the beads settle. Spin down briefly in a benchtop centrifuge after the second wash, resettle beads, and remove as much residual wash buffer as possible.
Dry the beads (caps open) until now much wash buffer is visible (5-10 minutes).
Elute sample in the following volumes depending on your chosen method of sequencing:
FOR PROMETHION: Resuspend sample in 33 µL REB elution buffer (also from ONT kit) and rotate for 10 min at RT. Reseat beads on the magnet, and move elution to a new tube.
FOR MinION: Resuspend sample in 13 µL REB elution buffer (also from ONT kit) and rotate for 10 min at RT. Reseat beads on the magnet, and move elution to a new tube.Elute sample in 13 µL REB elution buffer (also from ONT kit) with a 10 min incubation at RT.
Insert the flow cell into your sequencer and check its pores.
The next step (step 11) gives sequencing instructions for promethion flow cells. MinION instructions are in Appendix 5. Go there now if you are running a MinION.
Make the following priming mix and vortex. If you plan on running more flow cells than can sequence at once, only prep the mix for one batch at a time. You need 1000 µL per flow cell. Keep the mix on ice.
No need to make the master mix if you only after sequencing 1 flow cell.
Promethion Priming Mix Recipe
Mix 100 µL sequencing buffer (SB) with 68 µL library solution (LIS) for each sample.
Add 32 µL of library to each SB + LIS mix. Mix by pipetting and do not introduce bubbles.
You should already have equilibrated the flow cell to RT for 20 minutes and checked its pores during the RLA cleanup, but if you didn't, do that now.
If you are using a used flow cell, make sure the inlet port is CLOSED and remove all waste from either of the waste ports.
Open inlet port by rotating the valve / cover clockwise (it’s the thing with the screw).
Insert the pipet tip into the inlet port (the hole marked 1), and slowly turn the pipet wheel to draw back 20-30 µL, or until you see a small volume of yellow buffer entering the tip.
The amount you have to turn pipet may vary depending on brand of pipet tip and how well it is seated in the hole.
DO NOT REMOVE MORE THAN A FEW µLS OR YOU WILL EXPOSE THE PORES TO AIR.
Slowly load 500 µL of the priming mix into the inlet port (same hole marked 1) WITHOUT INTRODUCING AIR. CHECK TO MAKE SURE NO AIR IS AT THE TIP OF YOUR PIPET. IT IS BETTER TO USE A FEW µLs LESS THAN 500 THAN TO INTRODUCE AIR.
Close the rotating valve to avoid air getting into the port. If you forget, back out another small amount of liquid before adding the rest of the priming mix.
Wait 5 mins, then add another 500 µL. DO NOT INTRODUCE AIR.
Mix library by gentle pipetting just before loading. Do not introduce bubbles.
Insert a p200 tip into the inlet port and add slowly add 190 µL of library WITHOUT INTRODUCING AIR. CHECK TO MAKE SURE NO AIR IS AT THE TIP OF YOUR PIPET. IT IS BETTER TO USE A FEW µLs LESS THAN 500 THAN TO INTRODUCE AIR.
Rotate the valve to the closed position (below) to seal the inlet port.
Put the light sheild on. Place a sticker on the light sheild and label the flow cell with the library you just input.
Configure MinKNOW for RNA004 direct RNA sequencing with the following settings:
POD5 should be the selected output, 20 nts the minimum read size (200 nts may be the default), and do not filter reads.
You can now begin the sequencing run.
After your tRNA sequencing run has reached the desired number of reads, you can stop sequencing. Make sure you only tell it to stop sequencing NOT to stop sequencing and basecalling.
The flow cell can be removed before basecalling is finished, but a new flow cell can't be inserted into that position until basecalling is finished. If you do insert one, you won't be able to use that position until basecalling is complete.
Make the following wash mix. You need 400 µL per flow cell. If you'll be washing more cells later on, only make enough mix for the cells you plan to wash right away.
Close the inlet port. Remove all liquid from the waste port (labelled 2 or 3).
Open the inlet port. Pull out a SMALL amount of liquid to remove the air gap. DO NOT PULL OUT MORE THAN A FEW µLS OR YOU WILL EXPOSE THE PORES TO AIR.
Add 200 µL of the mix mix to the inlet port and close it.
Wait 5 minutes and add the remaining 200 µL. If you forget to close the port, reextract a small amount of liquid from the inlet port to remove any air before adding the remaining 200 µL wash mix.
Close the port and wait 1 hour for the DNase in the wash mix to work.
With the port closed, extract all the waste.
Open the port and remove a small amount of liquid from the inlet port.
Add 500 µL storage buffer to the flow cell. Close the port.
Go check how many pores are still active on the flow call. Write the number down on a sticker on the light sheild.
We consider any Promethion with less than 200 pores to be dead. Collect dead flow cells somewhere in your lab because ONT offers a free flow cell recycling program for dead cells.
Store at 4ºC until you want to use it again.
Mix 100 µM oligos at a 1:1 molar ratio in 10 mM Tris HCl and 50 mM NaCl (100 µL total). Add 1 µL RNase inhibitor.
Aliquot into 4 PCR tubes.
Heat the mix to 75ºC for 15 seconds and then cool to 25ºC at 0.1 ºC/second.
The final aliquots should be 10 µM.
This table is for you if when you nanodropped your small RNA fraction after deacylation, you got less than 30 ng/µL. If you got less than 10 ng/µL, I would caution you against proceeding at all since you may not get enough product at the end to sequence.
Add your ng/µL concentration of tRNAs to the yellow box. Your ligation reaction recipe will be calculated to maintain a 1:1 tRNA:splint adapter molar ratio.
The table will calculate the pmol tRNAs found in 10 µL of your sample (this should be all of it). It will then calculate the pmol of splint adapters necessary to add to the reaction and the volume this amounts to. It will also calculate the amount of water you need to bring the reaction up to its total volume.
Note: This table will only do math for one sample that is under 30 ng/µL post-small RNA cleanup. If you have multiple samples that are under this threshold, simply copy the table for each sample. Set up the respective reactions accordingly.
Set up the following reaction adding enzyme last.
If you have used this appendix to set up your splint adapter ligations, you MUST use Appendix 3 to set up your RTA ligations.
Splint Adapter Ligation Calculations
This table is for you if you used Appendix 2.
Add your ng/µL concentration of tRNAs ligated to splint adapters to the yellow box. Your ligation reaction recipe will be calculated to maintain a 3:1 tRNA:RTA molar ratio.
The table will calculate the pmol tRNAs found in the 9 remaining µL of your sample. It will then calculate the pmol of RTA necessary to add to the reaction and the volume this amounts to. It will also calculate the amount of water you need to bring the reaction up to its total volume.
Note: You will have to duplicate this table if you duplicated the Appendix 2 table.
Note: The RTA adapter concentration from ONT is 1.4 µM.
Set up the following reaction adding enzyme last.
RTA Adapter Ligation Math - NEB T4 DNA Ligase
RTA Adapter Ligation Math - Watchmaker T4 DNA Ligase
Make the following priming mix and vortex. If you plan on running batches of flow cells, only prep the mix for the ones you are about to run. You need 1000 µL per flow cell.
MinION Priming Mix Recipe
Mix 37.5 µL sequencing buffer (SB) with 25.5 µL library solution (LIS) for each sample.
Add 12 µL of library to each SB + LIS mix. Mix by pipetting and do not introduce bubbles.
You should already have equilibrated the flow cell to RT for 20 minutes and checked its pores during the RMX cleanup, but if you didn't do that now.
Insert the pipet tip into the priming port (beneath the rotation valve) and slowly turn the pipet wheel to draw back 20-30 µL, or until you see a small volume of yellow buffer entering the tip.
The amount you have to turn pipet may vary depending on brand of pipette tip and how well it is seated in the hole.
DO NOT REMOVE MORE THAN A FEW µLS OR YOU WILL EXPOSE THE PORES TO AIR.
Pipet 800 µL priming mix into the priming port WITHOUT INTRODUCING AIR. CHECK TO MAKE SURE NO AIR IS AT THE TIP OF YOUR PIPET. IT IS BETTER TO USE A FEW µLs LESS THAN INTRODUCE AIR.
Pipet the remaining 200 µL priming mix into the priming port WITHOUT INTRODUCING AIR.
Load 80 µL library and DO NOT INTRODUCE AIR. IT IS BETTER TO LEAVE BEHIND A FEW µLs THAN INTRODUCE AIR.
Put the light shield on. Label light shield with a sticker.
Configure MinKNOW for RNA004 direct RNA sequencing with the following settings:
POD5 should be the selected output, 20 nts the minimum read size (200 nts may be the default), and do not filter reads.
You can now begin the sequencing run.
After your tRNA sequencing run has reached the desired number of reads, you can stop sequencing. Make sure you only tell it to stop sequencing NOT to stop sequencing and basecalling.
The flow cell can be removed before basecalling is finished, but a new flow cell can't be inserted into that position until basecalling is finished. If you do insert one, you won't be able to use that position until basecalling is complete.
Make the following wash mix. You need 400 µL per flow cell. If you'll be washing more cells later on, only make enough mix for the cells you plan to wash right away.
Close the SpotON port and the priming port. Remove all liquid from the waste port (labelled 2 or 3).
Open the priming port. Pull out a SMALL amount of liquid to remove the air gap. DO NOT PULL OUT MORE THAN A FEW µLS OR YOU WILL EXPOSE THE PORES TO AIR.
Add 200 µL of the mix mix to the priming port and close it.
Wait 5 minutes and add the remaining 200 µL. If you forget to close the port, reextract a small amount of liquid from the inlet port to remove any air before adding the remaining 200 µL wash mix.
Close the port and wait 1 hour for the DNase in the wash mix to work.
With the port closed, extract all the waste.
Open the priming port and remove a small amount of liquid.
Add 500 µL storage buffer to priming port and close it.
Go check how many pores are still active on the flow call. Write the number down on a sticker on the light sheild.
Store at 4ºC until you want to use it again.
