PCR Barcoding Protocol

After FACS sorting and overnight outgrowth, colony PCRs with well-specific barcoded primers encode the plate and well position on each amplicon. PCR barcoding This enables multiplexed amplicon sequencing to determine the sequence identity of each well.

Overview

The barcoding strategy involves:

1. Well-specific primers containing unique DNA barcodes
2. PCR amplification of the variable region from bacterial lysate
3. Pooling of barcoded amplicons across all plates
4. Sequencing of the pooled library
5. Computational demultiplexing to assign reads to wells

Barcode Design

We recommend ordering primers described in Long et al. 2024 which have been optimized for multiplexed, long-read amplicon sequencing. The adaptor region can be designed to target a common promoter-terminator pair to sequence the entire gene on a single amplicon.

Materials

Item	Supplier	Catalog #
Taq DNA Polymerase	NEB	M0273S
Thermopol Buffer	NEB	B9004S
Barcoded LevSeq primers (384 wells)	IDT	Custom order
96-well PCR plates	Bio-Rad	HSP9601
PCR plate seals	Bio-Rad	MSB1001
Ampure XP beads	Beckman Coulter	A63880

Protocol

1. Prepare Primer Plates

1. Order barcoded primers at 100 µM stock concentration
2. Dilute to 10 µM working concentration in nuclease-free water
3. Aliquot into 384-well plates matching your plate layout:
   • Forward primer plate (contains universal reverse primer in all wells)
   • Reverse primer plate (contains well-specific barcodes)
4. Store at -20 °C

2. Colony PCR from 384-Well Plates

PCR setup (per well):

• 5 µL KAPA HiFi HotStart ReadyMix (2×)
• 0.25 µL forward primer (10 µM)
• 0.25 µL reverse primer with barcode (10 µM)
• 0.5 µL bacterial culture (direct from 384-well plate)
• 4 µL nuclease-free H₂O
• Total: 10 µL

1. Using a multichannel pipette, transfer 0.5 µL from each well of the 384-well plate to a PCR plate
2. Add master mix + primers to each well (can use liquid handler for automation)
3. Seal plates and centrifuge briefly
4. Run PCR program:
   • 95 °C, 3 min (initial denaturation + cell lysis)
   • 25× [98 °C, 20 s; 60 °C, 15 s; 72 °C, 30 s]
   • 72 °C, 2 min
   • 4 °C, hold

3. Verify Amplification

1. Run 2 µL from 8–12 random wells on a 1% agarose gel
2. Verify presence of expected band size
3. Check for non-specific amplification

4. Pool Barcoded Amplicons

1. Pool 2 µL from each well of all PCR plates into a single tube
2. Mix thoroughly by vortexing and inversion
3. Purify pooled amplicons using DNA Clean & Concentrator:
   • Follow manufacturer's protocol
   • Elute in 30–50 µL nuclease-free water
4. Quantify by Qubit or NanoDrop
5. Verify size distribution by TapeStation or Fragment Analyzer

File Formats

Barcode CSV

Create a CSV file mapping each well to its barcode sequence. This file is required for demultiplexing:

plate,well,barcode
plate_1,A01,ACGTACGTACGT
plate_1,A02,TGCATGCATGCA
plate_1,A03,GCTAGCTAGCTA
plate_1,A04,CGATCGATCGAT
plate_2,A01,AGTCAGTCAGTC
plate_2,A02,TCGATCGATCGA

Column definitions:

• plate: Unique identifier for each 384-well plate (e.g., plate_1, plate_2)
• well: Well position in standard format (A01–P24 for 384-well)
• barcode: DNA barcode sequence (typically 8–16 bp)

Quality Control

Expected Metrics

• PCR success rate: 95–98% of wells with growth should amplify
• Pooled concentration: 50–200 ng/µL after purification
• Fragment size: Insert + ~40 bp (primers + barcodes)

Troubleshooting

Problem	Possible Cause	Solution
No amplification	Insufficient template, primer issue	Increase template to 1 µL, check primer concentration
Multiple bands	Non-specific amplification	Optimize annealing temperature, redesign primers
Low yield after pooling	Many wells failed PCR	Check cell viability before PCR, optimize cycling
Unequal representation	Variable amplification efficiency	Normalize input or use fewer PCR cycles

Sequencing Submission

Submit the pooled, purified amplicon library for sequencing:

Plasmidsaurus (Recommended)

# Submit purified pool (500 ng - 2 µg in 15 µL)
# Turnaround: 3-5 business days
# Cost: ~$15 per sample (volume discounts available)

Oxford Nanopore (In-House)

# Use ligation sequencing kit (SQK-LSK109 or newer)
# Target: 200-500 ng input DNA
# Run time: 12-48 hours
# Cost: ~$500-900 per flow cell

Illumina MiSeq

# Submit 20-50 ng at 4 nM
# Requires PhiX spike-in (10%) for low-diversity libraries
# Run time: 24-56 hours
# Cost: $600-1500 per run

Next Steps

After receiving sequencing data, use the uSort-M CLI to demultiplex and analyze your results. LevSeq barcodes are built into the pipeline — no separate barcode CSV is needed.

usortm demux project/ \
  --fastq sequencing_data.fastq \
  --library-csv variants.csv

See Demultiplexing for detailed instructions.