DNA library preparation

CiBER-seq plasmid barcode library prep 

Introduction

This protocol is for preparing plasmid barcode sequencing libraries from yeast transformed with barcoded gRNA libraries including https://benchling.com/s/seq-e1JKDpvCq23IVfYOeAUf, https://benchling.com/s/seq-Ac30OQSFglXbXEJFYhuQ, https://benchling.com/s/seq-S5cQupYmmAS5YGu3hWZU or equivalent. These plasmid libaries were constructed with a T7 site for in-vitro transcription reactions and a 5nt identifier near the barcode to allow pooling of multiple yeast strains and/or libraries within a single experiment.

Materials

› Yeast Pellets, stored at -80 or flash frozen

› Zymo yeast miniprep kit (Zymo #D2004)

› T7 HiScribe Kit (NEB #E2040S)

› Suitable restriction enzyme (depending on plasmid library) to linearize the plasmid backbone, as long as it does not cut between the T7 site and the reverse transcription priming site. For libraries with barcodes in the 3'UTR of yeast citrine, MfeI works. For libraries with barcodes in the 3'UTR of mCherry, PvuII works.

› Protoscript II reverse transcription kit (NEB #M0368L)

› NEB-next dual indexing primers (NEB #E7600S)

› For barcoded citrine plasmid libraries use

  › Primer: 511 DNA_IVT_RT: GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT TGGTCCAGTCTTGTTACCAGACAACC

› For barcoded mCherry plasmid libraries use

  › Primer: 546 mcher IVT_RT: GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT TCAAGTTGGACATCACCTCCCAC

› For custom barcoded ORF plasmid libraries use

  › A custom designed primer with 5' adapter GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT (adds the compatible priming site for the i7 NEB #E7600S dual indexing primer) and a 3' end specific to a given ORF that will prime an RT reaction toward the random nucleotide barcode with roughly 150nt intervening sequence. (This ensures you'll have a final dual-indexed sequence that is ~300nt long, which is short enough to cluster well on the Illumina sequencer but long enough to do any necessary size selection to get rid of primers and primer dimers)

› RNAse A (ThermoFisher #EN0531)

› RNAse H (NEB #M0297S)

› Zymo DNA clean and concentrate kit (Zymo #D4013)

› Zymo RNA clean and concentrate kit (Zymo #R1017)

› Q5 polymerase (NEB #M0491L)

› Thermocycler

› Thermomixer

› Incubator

› Nanodrop or similar method for RNA concentration quantification

› Tape Station/Bioanalyzer + necessary Tape reagents

› AMPure XP beads or similar DNA size selection (if necessary, based on results of library prep)

Procedure

DNA library prep from yeast pellets

1. Thaw 50 mL of frozen pelleted yeast from the turbidostat:
Its good to harvest, pellet, and freeze down (at -80°C) extra yeast so that you have something to go back to in case something in the process fails. For reference, the yeast in the turbidostat should be OD roughly 0.8-1.2 Given that for haploid yeast 1mL of OD1 = 10^7 cells, 50 mL (~500 million cells) should be enough to cover a 240,000 barcoded guide library with about 2000x coverage. Note that you'll lose plasmid along the way in the yeast miniprep, so aiming for 25-50mL should still put you in a comfortable spot coverage-wise.

2. Follow the Zymo Yeast miniprep according to manufacturer's instructions for extracting plasmid from yeast liquid culture, with the following protocol adjustments. Resuspend the pellet in 1mL of solution 1/digestion buffer and add 30uL of zymolase. Triple the digestion time to ensure efficient plasmid extraction (3hours) and perform this with constant mixing (either in a thermomixer or shaking incubator). Scale up volumes of solution 2 and solution 3 by 5x to match the 5x scale of solution 1. Distribute across multiple eppendorfs so the entire reaction can be centrifuged and sequentially pass the supernatants across the same column. To maximize DNA recovery, perform the final plasmid elution using 20uL nuclease-free water warmed to 37°C.

3. Digest extracted plasmid (all of it) with MfeI or appropriate restriction enzyme to linearize the plasmid. I do 2-3 hour digestion to make sure complete digestion has occurred. Column clean according to Zymo clean and concentrate kit instructions. Eluting using 20uL nuclease-free water warmed to 37°C will maximize DNA recovery.

4. Perform an IVT reaction using the T7 HiScribe kit manufacturer's recommendations for small length product (30uL reaction, short transcripts, overnight at 37°C). I usually set this up in the 37°C incubator to prevent evaporation/condensation on the top of the lid.

5. Use the T7 HiScribe suggested DNase1 treatment to remove template DNA (add 2uL DNaseI from the kit and digest at 37C for 15 minutes). Column clean the RNA according to Zymo RNA clean and concentrate kit (Zymo #R1017) manufacturer instructions.

6. Quantify in-vitro transcribed RNA via nanodrop and reverse transcribe 2ug of IVT product according to protoscript II (NEB #M0368L) manufacturer’s protocol. (This will be a 2x scaled reverse transcription reaction, since the protocol recommends 1ug of RNA per reaction) Use the primer corresponding to your plasmid library (511 DNA_IVT_RT,546 mcher IVT_RT, or custom primer) to perform the RT reaction. This primer adds the i7 annealing site for downstream library prep. The plasmid already contains the i5 (TruSeq R1) priming sequence adjacent to the random nucleotide barcode.
As a side note, T7 has non-templated polymerase activity (especially in this situation where it is overnight and very low input) so only roughly 10% (estimated) is actually the IVT of the intended template. The rest is presumably randomly-polymerized RNA. These randomly-polymerized RNA species generally don't cause issues as they won't be reverse transcribed or PCR amplified in downstream steps.

7. Treat RT product for 1 hour at 37C with 0.5uL RNAse A and 0.5uL RNAse H. Then column clean DNA using ssDNA 7:1 ratio binding buffer according to kit instructions (Zymo #D4013).

8. The purified ssDNA now contains the two annealing sites compatible for use with NEB i5/i7 dual index primers. Assign primer pairs to each sample, making sure to choose a set of dual index primers that are compatible for pooled sequencing and will allow the samples to be separated by i5/i7 adapter sequence. Refer to the dual index primer manual for more information. Perform Q5 PCR amplification with 1/2 the column-cleaned product as template and NEB i5/i7 dual index primers according to manufacturer's instructions. Save the other 1/2 column- cleaned product to go back to if necessary. Use annealing temp 72C, and 10sec extension. Aim for low number of cycles, to avoid overamplification of PCR product (ideally 6-10, but may need to adjust depending on how the bioanalyzer trace looks).

9. Column clean the PCR product according to Zymo kit instructions and elute in nuclease-free water.

10. Run a 1:10 dilution of your samples on a Tape Station/Bioanalyzer to determine the concentration and sizing of each amplified barcode library. One sharp peak at the expected amplicon size, ~330bp (or other size if using custom reverse transcription primer), indicates a successfully-prepared sample. An additional peak that runs at a high molecular weight may indicate overamplification, as amplicons that melt and re-anneal form bubbles at the mismatched barcode sequence which tend to run slower on the Tape Station. I aim for this over-amplification peak to be <10% the molarity of the ~330bp peak. Over-amplified samples can be re-prepared from the remaining 1/2 column-cleaned reverse transcription product using fewer PCR cycles. Free primers and primer dimers in the sample should be removed using AMPure XP size selection or similar size selection strategy as they affect the quality of downstream sequencing.

11. Successfully-prepared samples can be pooled and submitted for HiSeq or NovaSeq

RNA library prep from yeast pellets

12. See "Dual Promoter CiBER-seq library prep" for directions on how to prepare expressed RNA barcode sequencing libraries.