Automated Bar-Seq Library Preparation and Pooling
Nina Alperovich, David Ross
Disclaimer
The protocol outlined in this document was created as a part of the Pooled, Growth-Based Assays for Protein Function Measurements pipeline for Align to Innovate’s Open Dataset Initiative. Align to Innovate is a non-profit research organization operating under open science principles with the goal of improving science research with programmable experiments. The Open Datasets Initiative is working to accelerate community-driven science with the use of automated labs to pioneer robust data collection methods and curated, high-fidelity, public biological datasets amenable to machine learning. This work was supported by Align to Innovate’s Open Datasets Initiative which receives philanthropic funding in part from Griffin Catalyst.
Abstract
Protocol for automated Bar-Seq Library preparation
This protocol prepares 96 DNA samples, representing 24 samples from 4 different timepoints, for multiplexed Illumina sequencing. The process starts with two rounds of PCR, each followed by a bead-based cleanup. The first round of PCR attaches primers that serve as tags to identify the timepoint and sample (e.g., time-points 1-4 in the Pooled, Growth-Based Assay Protocol). The second round of PCR attaches flow-cell adapters required for Illumina sequencing. Following the PCR and cleanup steps, the protocol outlines a procedure for pooling all samples together to ensure balanced representation during sequencing. This process starts with quantifying each sample's DNA concentration and then diluting and pooling together samples from the same timepoint. The pooled samples' concentrations are then measured, and a small quantity of each timepoint sample is run on a gel to ensure the cleanup process was successful. The final step is to dilute and pool of all timepoint samples into one multiplexed tube for subsequent Illumina sequencing.
NOTES:
- Before implementing this protocol, prepare the magnetic bead suspension following the Preparation of Sera-mag SpeedBeads protocol. Preparation of Sera-mag SpeedBeads protocol.
- This protocol should be implemented after the Automation Protocol for Plasmid DNA Extraction from E. coli protocol
- A fragment analyzer can also be used instead of a gel to determine if the PCR clean-up process was a success
Steps
Transfer Plasmid DNA to PCR Plate and Dilute with DI water
Pre-heat the on deck thermocycler (ODTC) for 1st PCR step
Remove lids from PCR-Plate 1, Sample-Plate, and Reagent Plate
Pipette 10uL nuclease-free water to each well in PCR plate 1
- Mix 6x after dispensing
- dispense 1 mm above bottom of well
- mix volume 25 uL
- mixing speed 75 uL/s
- mix position 0 mm (same as dispense position)
- liquid following OFF
Transfer 35uL of extracted plasmid DNA to each well of PCR Plate 1
- This step is done 24 samples (one time point) at a time, with user instructions to load each plasmid sample input plate (without lids)
- aspirate 1 mm below liquid surface, with liquid following ON
- dispense 0.5 mm below liquid surface, with liquid following ON
User instruction to remove plasmid sample input Sample-Plate
Run First PCR - using primers to identify samples from each timepoint
Add 28.125 uL Master Mix with reverse primer for the appropriate timepoint and sample to each corresponding well of PCR plate 1
- dispense 0.5 mm below liquid surface, with liquid following On
- no mixing
Add 28.125 uL Master Mix with forward primer for the appropriate timepoint and sample to each corresponding well of PCR plate 1
- Mix 10x after dispensing
- dispense 0.5 mm below liquid surface, with liquid following On
- mix volume 70 uL
- mixing speed 75 uL/s
- mix position 1 mm (below dispense position)
Put a PCR plate lid 1 on the PCR plate 1.
Move PCR-Plate 1 to ODTC
Run First PCR Rx using conditions (101.25 uL volume) using the following cycles:
- 98 C for 60 s
- 3 cycles at:
- 98 C for 10 s
- 58 C for 20 s
- 72 C for 20 s
-
72 C for 60 s
-
23 C for 10 s
First PCR Cleanup Part 1: Bind template plasmid DNA to the beads and keep the supernatant
Pipette 54 uL magnetic bead suspension into each well of Midi plate 1
- Bead ratio: 0.6x
- Mix the bead suspension 10x before pipetting
- Get new tips for each well
- Mix bead suspension again 2x before pipetting each well
- Pipette bead suspension into Midi plate during the Step 1 PCR
Move PCR plate 1 from ODTC; take lid off of PCR plate 1
Close ODTC; run PCR pre-heat method on ODTC (to get ready for Step 2 PCR)
Shake Midi plate 1 with bead suspension for 10 second at 1800 RMP
Transfer 90 uL of each sample from PCR plate 1 to Midi plate 1
Mix 10x after dispensing
- dispense 0.5 mm above bottom of well
- mix volume 108 uL
- mixing speed 100 uL/s
- mix position 2 mm (below liquid surface)
- liquid following ON
Incubate for 7 minutes at room temperature
Move Midi plate 1 to magnet base
Wait for 4 minutes
During 4 minute wait, move Midi Plate No. 2 to
HHS.
Transfer 134 uL of supernatant from Midi plate 1 to Midi plate 2
First PCR Cleanup Part 2: bind PCR product to beads and discard the supernatant
Pipette 80.4 uL bead suspension to Midi plate 2 (on HHS)
- Bead ratio: 0.6 for LacI
Mix the bead suspension 10x before pipetting
Get new tips for each well and mix bead suspension again 2x before aspiration for each well
Mix 10x after dispensing
- dispense 0 mm below liquid level
- 10 mix cycles
- mix volume 193 uL
- mixing speed: 100 uL/s
- mix position 1 mm (below liquid surface)
- liquid following ON
Incubate for 7 minutes at room temp, without shaking
During 7 minute wait, throw Midi Plate
No. 1 to waste
Move Midi plate 2 to magnet base
Wait for 5 minutes
Remove and discard most of the supernatant (209.4 uL)
Pipette 200 uL 80% ethanol into Midi plate 2 (on magnet)
- Jet dispense, from 20 mm height, using one tip for each row
Remove ethanol supernatant and discard
Using 50 uL tips, remove remaining supernatant and dispose
Move Midi plate 2 to HHS
Wait 30 s to allow ethanol to finish drying
Pipette 55 uL DI water into Midi plate 2
Mix 2x after dispensing
- dispense 1 mm above bottom of well
- mix volume 41.25 uL
- mixing speed 100 uL/s
- mix position 0 mm
- liquid following OFF
Shake at 1800 RPM for 10 s
Shake at 400 RPM for 2 minutes
Incubate for 3 minutes at room temp without shaking
Move Midi plate 2 to magnet base
Start timer for 5 min incubation
While waiting, start next step by preparing the 2nd PCR reaction
Run 2nd PCR: using primers for Illumina multiplexing
Prepare PCR plate 2 by pipetting 54 uL of 2x master mix (containing both forward and reverse universal primers for Illumina sequencing) into each well
Wait for 5 minute timer from the 1st PCR cleanup to finish.
Transfer 45 uL of supernatant from Midi plate 2 to PCR plate 2
Mix 10x after dispensing
- dispense 0.5 mm below liquid surface
- mix volume 69.3 uL
- mixing speed 100 uL/s
- mix position 1 mm (lower than dispense position)
- liquid following ON
Put a new PCR plate lid on PCR plate 2.
Move PCR-Plate 2 to ODTC
Run PCR plate 2 (99 µL) using conditions:
- 98 C for 60 s
- 15 Cycles at:
- 98 C for 10 s
- 72 C for 30 s
-
72 C for 120 s
-
10 C for 20 s
Note: during the 2nd PCR run, throw Midi plate 2 to waste and prepare Midi plate 3 following the instructions below.
Second PCR cleanup: Bind PCR product to beads and discard supernatant
Move Midi plate 3 to HHS (while 2nd PCR is running)
Pipette 89 uL bead suspension to Midi plate 3
- Bead ratio 1.0x
Mix the bead suspension 10x before pipetting
Get new tip for each well & mix beads suspension again 2x before pipetting each well
Wait for PCR to finish
Move PCR plate 2 from ODTC; take lid off of PCR plate 2
Shake Midi plate 3 with bead suspension for 10 second at 1800 RMP
Transfer 89 uL from PCR plate 2 to Midi plate 3
Aspirate 0.5 mm above bottom of well with liquid following OFF
Mix 10x after dispensing
- dispense 0.5 mm above bottom of well
- mix volume 133.5 uL
- mixing speed 100 uL/s
- mix position 2 mm (below dispense position)
- liquid following ON
Incubate for 7 minutes
Move Midi plate 3 to magnet base
Wait for 5 minutes
Remove and discard most of the supernatant (173 uL)
Pipette 200 uL 80% ethanol into Midi plate 4
- jet dispense, from 20 mm height, using one tip for each row
Wait 30 s
Remove ethanol supernatant and discard
Using 50 uL tips, remove remaining supernatant and dispose
Move Midi plate 3 to HHS
Wait 30 s to allow ethanol to finish drying
Pipette 50 uL Elution buffer into Midi plate 3
Mix 2x after dispensing
- dispense 1 mm above bottom of well
- mix volume 37.5 uL
- mix position 0 mm
- liquid following OFF
Shake at 1800 RPM for 10 s
Shake at 400 RPM for 2 minutes
Incubate for 3 minutes without shaking
Move Midi plate 3 to magnet base
Wait for 5 minutes
- Remove lid from Output plate during wait
Transfer 45 uL supernatant from Midi plate 3 to Output plate
Replace lid on Output plate
Pooling for Sequencing - Measure each sample's input plasmid DNA Concentration
Measure the DNA concentrations (Cn) for all input plasmid samples.
- Note: The expected concentration (Cn's) is typically between 5 ng/uL and 40 ng/uL, though some samples for time point 4, the final growth plate, may end up lower than that.
Quantify the amount of amplified DNA
The amount of amplified DNA should also be measured for each sample at each time point using Qubit or an equivalent DNA quantitative measurement.
- Expected yields are between 10 ng/uL and 30 ng/uL (in 45 uL), though some samples could have as little as 1 ng/uL (e.g., samples grown with the highest antibiotic concentration at the later time points).
Within each time point's samples, calculate the samples' final pooling volumes (Vf)
Within each time point's samples (i.e., the 24 samples coming from each growth plate), identify the the four lowest-concentration samples.
Calculate the mean of their concentrations in ng/µL, and define this concentration as the low concentration limit (Clow ) for that time point.
For each sample, use the DNA concentration in ng/µL (Cn) and the following equation to calculate the pooling volume (Vn) for each sample:
- Vn = Clow/(Cn + 0.1 ng/uL)*15 µL
For each sample, set the final pooling volume (Vf) based on the following conditions:
- If Vn< 4 µL, set Vf = 4 µL
- If Vn> 15 µL, set Vf = 15 µL.
Within each time point, pool all samples and measure the pooled-DNA concentration
Within each time point's samples (i.e., the 24 samples coming each growth plate), pool together all samples into one container according to their final pooling volumes (Vf's) and store the remaining un-pooled samples at -20C.
- Note: This will give one pooled sample for each time-point (four pooled samples in total), each at a concentration of approximately 20 ng/uL and with a volume of 100 to 300 uL. This is much more DNA than is needed for sequencing.
For each of the four pooled samples, re-measure the DNA concentration and then store them at -20C until the next step can be accomplished.
Run a gel for each time point
Run a gel for each time point, and compared these results with those obtained during BarSeq method development.
- The gels should not have any visible bands that are shorter than the expected amplicon size (315 bp)
- Shorter bands indicate incomplete PCR cleanup that could result in a very low sequence count from Illumina sequencing.
- If the gels results look correct, proceed to the next step
Dilute each time point's sample and pool all samples for sequencing
To prepare samples for the sequencing provider, follow their instructions for pre-made Illumina libraries.
- Note: They usually give the sample requirements in molar concentration, typically 5 nmol/L, which is about 1 ng/uL for the BarSeq libraries. They usually require a minimum of 20 uL to 30 uL per lane of sequencing.
- As an example, using the numbers above, a sample protocol might be to dilute 5 µL of each pooled sample into nuclease-free water to a concentration of 1 ng/uL. This will give much more than the minimum required volume, but it avoids the extra variability from pipetting smaller volumes.
Combine the diluted time point samples to be run on the sequencer. In the following manner:
- For the first run: start with a single lane of NovaSeq, with all four time points combined in equal proportions. Then, after analyzing that data, if you need more data, order a second lane and send a second sample with only time points 2-4.
- For consecutive runs: order a single lane of NovaSeq with the combine the time points with the following ratios: ⅓:1:1:1 for time points 1-4.
