Sep 30, 2024
PCR amplification of the barcode region
- Melinda Wheelock1,
- Nicholas Popp1,
- Raining Wang1,
- Douglas Fowler1
- 1University of Washington
Protocol Citation: Melinda Wheelock, Nicholas Popp, Raining Wang, Douglas Fowler 2024. PCR amplification of the barcode region. protocols.io https://dx.doi.org/10.17504/protocols.io.3byl49jjogo5/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: May 09, 2024
Last Modified: September 30, 2024
Protocol Integer ID: 99789
Funders Acknowledgement:
NHGRI (IGVF)
Grant ID: HG)11969
Abstract
This protocol details the gDNA amplification and plasmid DNA amplification.
Guidelines
This protocol will be written as if running a single sample through. Calculations will need to be made for any optimizations
- Annealing temperature, extension time, annealing time, and DNA concentration are all parameters to be optimized for each PCR reaction
- Optimizations are best done at 10-20 mL and run on a 2% agarose gel
This protocol is written for either gDNA amplification or plasmid DNA amplification.
- Differences will be noted in bold
Materials
Materials
● Micropipettes and tips: P2, P20, P200, P1000
● 0.2 mL 8-strip cap PCR tubes (clear) with lids
● 0.2 mL 8-strip cap PCR tubes (white) with lids (flat, clear)
● Primers
● SYBR Green I
● Q5 2x Master Mix
● Nuclease free water
● 70% ethanol (freshly made)
● AmPure XP magnetic beads
● Magnetic plate (for 0.2 mL PCR tubes or for 1.7 mL Eppendorf tubes)
● 1.7 mL Eppendorf tubes
● Agarose
● SYBR Safe (10,000x)
● 100 mL glass beaker
● 1x TBE (from carboy)
● 1kb+ DNA ladder
● 6x orange dye
● New straight-edge razor
● Freeze n Squeeze columns
● Lo-bind DNA tubes
● Qubit dsDNA HS Assay kit
● Qubit assay tubes
Equipment
● PCR thermocycler
● qPCR thermocycler (MiniOpticon)
● GelDoc
● Blue light gel box
● Orange protective eyewear
● Microwave
● Gel casting tray and gaskets
● Agarose gel electrophoresis box and power supply
● Benchtop microcentrifuge
PCR 1: Adding adapters and/or crossing the recombination junction
For gDNA, set up 8 identical PCR reactions as follows:
| A | B | C | D | |
| Reagent | Volume for 50 uL reaction | Volume for 8 reactions | Final concentration | |
| 10 mM forward primer (NP0492) | 2.5 uL | 20 uL | 0.5 mM | |
| 10 mM reverse primer (NP0546) | 2.5 uL | 20 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 200 uL | 1x | |
| gDNA template | 2500 ng (varies) | varies | 50 ng/mL | |
| Nuclease free water | varies | varies | n/a |
For plasmid DNA, set up 1 reaction as follows:
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (NP0492) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (NP0493) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| Plasmid DNA template | 10 ng (varies) | 0.2 ng/uL | |
| Nuclease free water | varies | n/a |
PCR 2: Nested PCR to add indices and cluster generators
For gDNA: set up 1 PCR reaction as follows in white PCR tubes with clear caps.
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (e.g. JS1000) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (e.g. NP0547) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| gDNA template | 8 uL (40% of elution) | varies | |
| 100x SYBR Green I | 0.25 uL | 0.5x | |
| Nuclease free water | 11.75 uL | n/a |
For plasmid DNA: set up 1 PCR reaction as follows in white PCR tubes with clear caps.
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (e.g. JS1000) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (e.g. JS391) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| Plasmid DNA template | 15 uL | varies | |
| 100x SYBR Green I | 0.25 uL | 0.5x | |
| Nuclease free water | 4.75 uL | n/a |
Before start
Notes about PCR 1:
The first PCR in the gDNA amplification process is designed to both add adapter sequences to one end of the amplicon as well as cross the recombination junction.
- The design of these primers should be such that the forward primer sits within the recombining plasmid and has a standard adapter sequence attached.
- The reverse primer should sit within the landing pad (usually at the start of BFP).
- The use of these two primers, in these two locations, ensures that the only gDNA that gets amplified is gDNA from cells that have successfully recombined.
○ Unrecombined cells will not be able to bind the forward primer, and leftover plasmid will not bind the second primer.
For this step, we run 8 identical reactions for two reasons:
- To reduce “jackpotting” where a random polymerase error gets propagated through the sequencing run.
- To ensure we use enough gDNA to cover our library, since gDNA is extremely large (>6 billion bp) and thus has very few copies even in thousands of ng of gDNA.
The process for plasmid DNA is identical, but we use 10 ng plasmid DNA instead of 2500 ng gDNA in the reaction and we only do 1 reaction instead of 8.
- The reverse primer for plasmid DNA, since it does not need to span a recombination junction, uses a different reverse primer that contains an adapter sequence (just like the forward primer) – use NP0493.
Notes about AmPure XP cleanup:
The amount of AmPure XP beads to use is dependent on the size of the amplicon that is being purified. Generally, AmPure XP cleans up smaller fragments, leaving larger ones to be eluted and saved.
- More beads allows smaller fragments to be retained.
Notes about PCR 2:
The second PCR in the gDNA amplification process is designed to reduce the size of the final amplicon to one that will cluster well on Illumina (smaller amplicons perform better) as well as add the sample indices and cluster generator sequences that will be used by Illumina to sequence and demultiplex samples.
- Each individual sample will require a separate pair of indices, one on the forward and one on the reverse primer.
○ We usually have 14 distinct indices for the forward primer and 14 distinct indices for the reverse primer.
- The indexes can be combined to create up to 196 unique index combinations (14 forward x 14 reverse) to multiplex up to 196 samples on the same Illumina run.
- In practice, we usually use less than 20 on a single Illumina run, which is entirely dependent on how many sequencing reads we need per sample compared to the number of sequencing reads produced by the Illumina kit.
○ The reverse primer will bind directly to the amplicon sequence and will contain a unique index and a cluster generator sequence (P7).
- This primer binds 5’ to the reverse primer in PCR1, since we need to reduce the size of the amplicon, usually to about 250-350 bases, for good Illumina sequencing.
- If the location of the barcode changes for whichever reason, it’s necessary to change these primers to match that new location.
○ The forward primer will bind to the adapter sequence added in PCR1 and will contain a unique index and the other cluster generator sequence (P5).
- The adapter sequence is designed such that we do not get amplification of any gDNA other than that which was amplified in PCR1. This ensures that we continue to not amplify random plasmid or gDNA that does not contain our barcoded region.
- For plasmid DNA, you will need to alter the reverse primer, since it has an adapter sequence to bind to.
PCR 1: Adding adapters and/or crossing the recombination junction
PCR 1: Adding adapters and/or crossing the recombination junction
For gDNA, set up 8 identical PCR reactions as follows:
| A | B | C | D | |
| Reagent | Volume for 50 uL reaction | Volume for 8 reactions | Final concentration | |
| 10 mM forward primer (NP0492) | 2.5 uL | 20 uL | 0.5 mM | |
| 10 mM reverse primer (NP0546) | 2.5 uL | 20 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 200 uL | 1x | |
| gDNA template | 2500 ng (varies) | varies | 50 ng/mL | |
| Nuclease free water | varies | varies | n/a |
For plasmid DNA, set up 1 reaction as follows:
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (NP0492) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (NP0493) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| Plasmid DNA template | 10 ng (varies) | 0.2 ng/uL | |
| Nuclease free water | varies | n/a |
Run 5 cycles of PCR on a standard thermocycler with the following parameters:
| A | B | C | |
| Step | Temperature | Time | |
| 1 | 98°C | 30 s | |
| 2 | 98°C | 10 s | |
| 3 | 61°C | 30 s | |
| 4 | 72°C | 30 s (1 min/kb) | |
| Go to 2 | 4x | ||
| 5 | 72°C | 2 min | |
| 6 | 4°C | hold |
AmPure XP cleanup
AmPure XP cleanup
13m
13m
Pool all 8 PCR reactions into a single 1.7 mL Eppendorf tube
For gDNA, add 0.8 mL AmPure XP beads for every 1 mL of product and mix by pipetting.
- Approximately 320 µL of beads to 400 µL of product, not accounting for evaporation.
For plasmid DNA, add 1 mL AmPure XP beads for every 1 mL of product.
- Approximately 50 µL of beads to 50 µL of product, not accounting for evaporation.
Incubate at Room temperature for 00:05:00 .
5m
Add tube to magnetic stand and let sit for 00:02:00 to pull beads to the side.
2m
Carefully aspirate with a pipette without disturbing the beads.
Wash with 2x volume of 70% ethanol.
Note
Approximately 800 µL of 70% ethanol.
Incubate at Room temperature for 00:02:00 on magnetic plate.
2m
Carefully aspirate with a pipette without disturbing the beads.
Repeat wash with 2x volume of 70% ethanol.
Allow any leftover ethanol to fully evaporate by leaving the lid off.
For gDNA: Add 21 µL nuclease free water to the beads and mix by vortex or pipetting.
For plasmid DNA: add 16 µL nuclease free water to the beads and mix by vortex or pipetting.
Incubate at Room temperature for 00:02:00 .
2m
Incubate at Room temperature for 00:02:00 on magnetic plate.
2m
Transfer the eluted DNA (~20 µL) to a new PCR tube.
PCR 2: Nested PCR to add indices and cluster generators
PCR 2: Nested PCR to add indices and cluster generators
For gDNA: set up 1 PCR reaction as follows in white PCR tubes with clear caps.
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (e.g. JS1000) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (e.g. NP0547) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| gDNA template | 8 uL (40% of elution) | varies | |
| 100x SYBR Green I | 0.25 uL | 0.5x | |
| Nuclease free water | 11.75 uL | n/a |
For plasmid DNA: set up 1 PCR reaction as follows in white PCR tubes with clear caps.
| A | B | C | |
| Reagent | Volume for 50 uL reaction | Final concentration | |
| 10 mM forward primer (e.g. JS1000) | 2.5 uL | 0.5 mM | |
| 10 mM reverse primer (e.g. JS391) | 2.5 uL | 0.5 mM | |
| 2x Q5 master mix | 25 uL | 1x | |
| Plasmid DNA template | 15 uL | varies | |
| 100x SYBR Green I | 0.25 uL | 0.5x | |
| Nuclease free water | 4.75 uL | n/a |
For gDNA: run 15-20 cycles of qPCR on the MiniOpticon or CFX with the following parameters:
| A | B | C | |
| Step | Temperature | Time | |
| 1 | 98°C | 30 s | |
| 2 | 98°C | 10 s | |
| 3 | 65°C | 30 s | |
| 4 | 72°C | 30 s (1 min/kb) | |
| Go to 2 | 19x | ||
| 5 | 72°C | 2 min | |
| 6 | 4°C | hold |
For plasmid DNA: run 9-12 cycles of qPCR instead.
| A | B | C | |
| Step | Temperature | Time | |
| 1 | 98°C | 30 s | |
| 2 | 98°C | 10 s | |
| 3 | 68°C | 30 s | |
| 4 | 72°C | 30 s (1 min/kb) | |
| Go to 2 | 19-24x | ||
| 5 | 72°C | 2 min | |
| 6 | 4°C | hold |
Note
IMPORTANT: stop the qPCR reaction when the relative fluorescence units (RFU) for all samples has risen to or above 0.3 (MiniOpticon) or 3000 (CFX). It does not need to complete the final extension or hold steps
- For gDNA: this is often around 15-20 cycles.
- For plasmid DNA: this is often around 9-12 cycles.
Gel extraction of amplicon
Gel extraction of amplicon
1h 17m
1h 17m
Make a 2% agarose gel in 1x TBE.
a. Melt in microwave at 30% power for 00:03:00 -00:04:00 , watch for over-boiling.
b. Mix in 12 µL SYBR Safe by swirling gently.
c. Pour into gel casting tray (make sure you have the gaskets in tightly!) after ensuring that the gel is level.
d. Insert a 10 well thick comb.
e. Cover with foil to protect from light until solidified.
7m
Fill gel box to maximum level with 1x TBE.
Insert gel (remove gaskets but keep plastic tray) into gel box vertically.
When adding DNA to the gel, skip every other lane to ensure less bleedover between samples.
Add 20 µL 1kb+ ladder to the first well.
Add all 50 µL of qPCR product to 10 uL 6x orange dye and load the entire volume into the gel.
Add 20 µL 1kb+ ladder to the last well.
Run the gel at 120V for 01:00:00 .
1h
Visualize gel on a blue light gel box.
Note
Wear orange safety glasses!
Using a brand new straight razor, carefully cut out the band of expected size.
Trim the edges of the gel so only illuminated sample is present in the final gel piece.
Note
Rotate in all directions to make sure.
Discard any extra gel, leaving the extracted piece with amplicon on the illuminated gel box.
Note
If running multiple samples to extract, move the gel to the side/back into the gel tray until the next step is complete.
Finely chop the gel containing the amplicon using the razor. Add the cubes to a labeled freeze n squeeze column.
Note
If running multiple samples, repeat these steps to cut out each band and place in a new freeze n squeeze column.
Place freeze n squeeze column(s) into -20 °C freezer for 00:05:00 .
5m
Spin sample at maximum speed in the tabletop centrifuge (>13.000 x g, 00:05:00 ).
5m
Collect DNA amplicon(s) from collection tube(s) and place in lo-bind DNA tube(s).
Qubit amplicon to determine DNA concentration
Qubit amplicon to determine DNA concentration
2m
2m
Collect one Qubit assay tube (0.5 mL) for each sample plus two tubes for standard DNAs.
- Label the lids, not the sides. Qubit measures through the sides of the tubes.
Make working solution by diluting dsDNA HS reagent 1:200 in dsDNA HS buffer in plastic tubes and mix by vortexing.
This comes out to 1 µL dsDNA HS reagent for every 199 uL dsDNA HS buffer.
You’ll need 200 µL for each sample + 400 µL for standards and pipetting error.
- For example, if you have 2 samples, you’ll need to make 800 µL of working solution (2x200 uL + 400 uL) by mixing 8 µL of dsDNA HS reagent to 792 µL dsDNA HS buffer.
Add 190 µL working solution to each of the two standards tubes.
Add 10 µL of each standard to the appropriate tube and mix by vortexing.
Add 198 µL working solution to each sample tube.
Add 2 µL of sample to the appropriate tube and mix by vortexing.
Incubate samples and standards at Room temperature for 00:02:00 .
2m
Take samples to Qubit reader (2.0).
- Ensure Qubit is set to DNA – dsDNA high sensitivity.
On the standards screen, select yes to read standards.
Read standard #1 by placing into reader, closing lid, and pressing read.
Remove standard #1 and repeat with standard #2.
Once the sample screen appears (should appear automatically), remove standard #2 and place in sample.
Close lid and press read.
- Record the concentration reported by Qubit (in ng/mL).
Repeat with each sample until all samples are read.
Back-calculate the actual sample concentration(s)
- ng/mL
- Qubit measures the concentration without accounting for the dilution into working solution.
Then, calculate the nM concentration of the amplicon.
Estimate the dsDNA molecular weight using this tool by copying the amplicon sequence from benchling: https://www.bioinformatics.org/sms2/dna_mw.html
- Make sure you have it selected as double stranded linear DNA
- Since the amplicon has N nucleotides, it will give you a range of values. Take the average of these two numbers
Use “molarity from mass and volume” from this tool to calculate a molar concentration: https://www.graphpad.com/quickcalcs/Molarityform.cfm
- Another option is to use the following equation if this tool no longer works for whatever reason: where [sample] is the sample concentration in ng/mL and MW is the molecular weight from the previous step.
Store DNA amplicon at -20 °C until ready for use.
- Requantify using Qubit at the time of use to account for evaporation or binding to tubes.
Sequencing
Sequencing
Pool the prepared libraries by molarity in proportions appropriate for the sequencing kit.
Sequence using an appropriate Illumina NGS sequencing kit.