Feb 07, 2023
Version 2
Cost-effective targeted nanopore sequencing of P. falciparum malaria V.2
This protocol is a draft, published without a DOI.
- Mariateresa de Cesare1,
- Mulenga Mwenda2,
- Anna E. Jeffreys1,
- Daniel J Bridges2,
- Jason A Hendry1
- 1Wellcome Centre for Human Genetics, Oxford;
- 2PATH
Protocol Citation: Mariateresa de Cesare, Mulenga Mwenda, Anna E. Jeffreys, Daniel J Bridges, Jason A Hendry 2023. Cost-effective targeted nanopore sequencing of P. falciparum malaria. protocols.io https://protocols.io/view/cost-effective-targeted-nanopore-sequencing-of-p-f-cnynvfveVersion created by NOMADS TEAM
Manuscript citation:
https://www.biorxiv.org/content/10.1101/2023.02.06.527333v1
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: February 07, 2023
Last Modified: February 07, 2023
Protocol Integer ID: 76526
Keywords: malaria, nanopore, amplicon
Funders Acknowledgement:
Bill and Melinda Gates Foundation
Grant ID: INV-003660
Abstract
This protocol outlines a cost-effective approach for amplicon sequencing of P. falciparum malaria from dried blood spots (DBS). The protocol can be conducted in ~2-3 days and costs ~USD $25/sample. We currently have two target panels, NOMADS8 and NOMADS16, which were developed using multiply (https://github.com/JasonAHendry/multiply).
Here, we describe how to generate the primer pools and prepare the library for sequencing. The protocol starts from dried-blood spot (DBS) extracted DNA. To date, most of our tests have been with samples extracted using Qiagen QIAamp kits (see Materials) and we observe good performance for samples with parasitemia > 1000p/uL. The assay performs better on fresher samples compared to those that have spent a long time as DBS. It is advisable to include positive controls in your experiment (for example, lab strains mixed with human DNA to mimic DBS-extracted DNA) as well as non-template controls.
We welcome your questions and/or feedback on the protocol.
References
- The sWGA protocol we use is modified from Samuel O. Oyola et al. (2016) Mal J.
- Lower extension temperature PCR inspired by Xin-zhuan Su et al. (1996) NAR.
- The one-pot sample barcoding protocol is modified from Josh Quick's One-pot native barcoding of amplicons
Materials
Major reagents
| Step | Item | Supplier | Item Code | |
| General | Qubit™ dsDNA HS Assay Kit (500) | ThermoFisher | Q32854 | |
| DNA Extraction | QIAamp DNA Micro Kit (50) | Qiagen | 56304 | |
| sWGA | phi29 DNA Polymerase | NEB | M0269L | |
| sWGA | 1000uM sWGA Primers | |||
| sWGA | NEB 10mM dNTP Solution Mix | NEB | N0447L | |
| Multiplex PCR | NOMADS16 Primers | |||
| Multiplex PCR | KAPA HIFI + dNTPS (100U) | Roche | KK2101 | |
| Barcoding | Native Barcoding Expansion 96 | ONT | EXP-NBD196 | |
| Barcoding | NEBNext Ultra II DNA Library Prep Kit with Sample Purification Beads* | NEB | E7103L | |
| Adapter Ligation | Ligation Sequencing Kit | ONT | SQK-LSK109 | |
| Adapter Ligation | NEBNext Quick Ligation Module | NEB | E6056L | |
| Sequencing | R9.4.1 Flow Cell | ONT | FLO-MIN106D |
Note that plasticware and laboratory basics like ethanol and nuclease-free water are not included here.
*This kit contains enough material perform sample barcoding for 480 samples, as well as SPRI beads.
sWGA Primer Pool
| Primer | Sequence | Quantity | Formulation | |
| Pf1 | ATATATATAT*A | 250 nmole | STD | |
| Pf2 | TATATATATAT*T | 250 nmole | STD | |
| Pf3 | TATATATATA*A | 250 nmole | STD | |
| Pf4 | TAATATATA*T | 250 nmole | STD | |
| Pf5 | TATATATATT*T | 250 nmole | STD | |
| Pf6 | ATTATTATTA*T | 250 nmole | STD | |
| Pf7 | TAATAATAAT*A | 250 nmole | STD | |
| Pf8 | AAAAAAAAAAA*A | 250 nmole | STD | |
| Pf9 | AATAATAATA*A | 250 nmole | STD | |
| Pf10 | TATTATATA*T | 250 nmole | STD |
The asteriks (*) indicates that the 3’ nucleotide is joined thorugh a phosphorothioate bond to inhibit the exonucleotide acitivity of phi29; note there is an extra charge per primer for the modification.
NOMADS8 Primer Pool
| Primer | Sequence | Quantity | Formulation | |
| CRT1_s8_F | TTTCATTGTCTTCCACATATATGACA | 25 nmole | STD | |
| CRT1_s8_R | TGCTCCTTTGACACAGTATGA | 25 nmole | STD | |
| DHFR_s0_F | AGCCATTTTTGTATTCCCAAATAGC | 25 nmole | STD | |
| DHFR_s0_R | ACCTCCTTAACAGAACTAGCCG | 25 nmole | STD | |
| DHPS_s14_F | AGAACAGCTATCGTATGAGGTACA | 25 nmole | STD | |
| DHPS_s14_R | TCACATGTTTGCACTTTCCTTT | 25 nmole | STD | |
| K13_s14_F | ATCCAAGCCTAGAACCTAATCC | 25 nmole | STD | |
| K13_s14_R | GGCGTAAATATTCGTGTTATAATTTCT | 25 nmole | STD | |
| MDR1part_s6_F | AGAGTTGAACAAAAAGAGTACCGC | 25 nmole | STD | |
| MDR1part_s6_R | CCAATGTTGCATCTTCTCTTCCA | 25 nmole | STD | |
| MSP2_s6_F | CACATGGACAACCTGCATCAC | 25 nmole | STD | |
| MSP2_s6_R | GTGCCACATTATAGCGCCCT | 25 nmole | STD | |
| PMI_s9_F | AAACAGACAATACCCGTCTACA | 25 nmole | STD | |
| PMI_s9_R | ACAAACTAGCCATTTGTATGCCT | 25 nmole | STD | |
| PMIII_s6_F | AAAAACGGGTCTGCCAAAAT | 25 nmole | STD | |
| PMIII_s6_R | TCAACCCAGCATGTGTTTCT | 25 nmole | STD |
NOMADS16 Primer Pool
| Primer | Sequence | Quantity | Formulation | |
| AMA1_S11_F | CACCTTGTTTTGAAACCTTTACACA | 25 nmole | STD | |
| AMA1_S11_R | TGTTGTACACATTTTCCCATAACA | 25 nmole | STD | |
| CSP_S3_F | CTTTTCCGGGGTCCCTTTCA | 25 nmole | STD | |
| CSP_S3_R | CGTGTAAAAATAAGTAGAAACCACGT | 25 nmole | STD | |
| HRP2_S12_F | TCGCTATCCCATAAATTACAAAACA | 25 nmole | STD | |
| HRP2_S12_R | ACCATAATAAATTACTTTCCAGTCACA | 25 nmole | STD | |
| HRP2dwn_S5_F | GCAGTAACATCATGGTTTTACATTACA | 25 nmole | STD | |
| HRP2dwn_S5_R | GGCTAATTTTTCAGGAGCTTCTACC | 25 nmole | STD | |
| HRP2up_S11_F | TGTGGGAATTTATAAGTTTCCAAGT | 25 nmole | STD | |
| HRP2up_S11_R | ACCACACCTACGTCAGTAACA | 25 nmole | STD | |
| HRP3_S12_F | AGCATATTATCTTCACTCACCCC | 25 nmole | STD | |
| HRP3_S12_R | GTGGTATTCAGAACATTCGGATTTATT | 25 nmole | STD | |
| HRP3dwn_S1_F | GCTAGGTGTATACGCACAGA | 25 nmole | STD | |
| HRP3dwn_S1_R | ATTCCCATGTTAAGCCTCTATTTTT | 25 nmole | STD | |
| HRP3up_S6_F | TCAATCTGAGGGATTAGTTGATGA | 25 nmole | STD | |
| HRP3up_S6_R | ACACCGTCAAATCCTCGATA | 25 nmole | STD |
NB: These primers are combined in addition to the NOMADS8 primers to create the full NOMADS16 panel.
Preparation of primer pools.
Preparation of primer pools.
Prepare the sWGA primer pool (see Materials).
If you have ordered the primers lyophilised, make them up to 1000uM in nuclease-free water.
To make the 1000uM sWGA primer pool, combine equal volumes of each primer (e.g. Pf1-Pf10) in a 1.5mL LoBind tube.
Store the primers at 4C.
Prepare the NOMADS8 or NOMADS16 primer pools (see Materials).
If you have ordered the primers lyophilised, make them up to 100uM in nuclease-free water.
If you want to run the NOMADS8 panel, use the table below to add the indicated volume of each 100uM primer stock to the primer pool. Afterwards dilute the primer pool to 10uM in nuclease-free water.
| F primer | R primer | Amplicon Size (bp) | Vol. into pool (uL)* | |
| CRT1_s8_F | CRT1_s8_R | 3874 | 6 | |
| DHFR_s0_F | DHFR_s0_R | 3463 | 3 | |
| DHPS_s14_F | DHPS_s14_R | 3656 | 3 | |
| K13_s14_F | K13_s14_R | 3826 | 1.5 | |
| MDR1part_s6_F | MDR1part_s6_R | 3773 | 1.5 | |
| MSP2_s6_F | MSP2_s6_R | 3720 | 3 | |
| PMI_s9_F | PMI_s9_R | 3101 | 6 | |
| PMIII_s6_F | PMIII_s6_R | 3468 | 3 | |
| TOTAL (uL) | 54 |
*This is the volume of each of the F and R primer to add. So for CRT1_s8; you will be adding 6uL of CRT1_s8_F and 6uL of CRT1_s8_R.
Note
For reasons of consistency, it is often advisable to make a larger volume pool than indicated above and split into aliquots.
Note
Why aren't all the volumes the same? In preliminary experiments we found that mdr1 and kelch13 amplicons were very abundant, whereas crt1 amplicons were relatively few. Thus the primer volumes were adjusted to mitigate against this imbalance.
If you want to run the NOMADS16 panel, use the table below to add the indicated volume of each 100uM primer stock to the primer pool. Afterwards dilute the primer pool to 10uM in nuclease-free water.
| F primer | R primer | Amplicon Size (bp) | Vol. into pool (uL)* | |
| CRT1_s8_F | CRT1_s8_R | 3874 | 6 | |
| DHFR_s0_F | DHFR_s0_R | 3463 | 6 | |
| DHPS_s14_F | DHPS_s14_R | 3656 | 3 | |
| K13_s14_F | K13_s14_R | 3826 | 3 | |
| MDR1part_s6_F | MDR1part_s6_R | 3773 | 1.5 | |
| MSP2_s6_F | MSP2_s6_R | 3720 | 3 | |
| PMI_s9_F | PMI_s9_R | 3101 | 3 | |
| PMIII_s6_F | PMIII_s6_R | 3468 | 3 | |
| AMA1_S11_F | AMA1_S11_R | 3138 | 1.5 | |
| CSP_S3_F | CSP_S3_R | 3107 | 3 | |
| HRP2_S12_F | HRP2_S12_R | 3097 | 3 | |
| HRP2dwn_S5_F | HRP2dwn_S5_R | 3333 | 3 | |
| HRP2up_S11_F | HRP2up_S11_R | 3366 | 3 | |
| HRP3_S12_F | HRP3_S12_R | 3444 | 3 | |
| HRP3dwn_S1_F | HRP3dwn_S1_R | 3304 | 3 | |
| HRP3up_S6_F | HRP3up_S6_R | 3066 | 6 | |
| TOTAL (uL) | 108 |
*This is the volume of each of the F and R primer to add. So for CRT1_s8; you will be adding 6uL of CRT1_s8_F and 6uL of CRT1_s8_R.
Note
Treat these volumes as suggestions. For the NOMADS16 panel, we did one round of amplicon balance optimisation but the balance can still be improved. If you find a combination of primer volumes that produces more balanced coverage across targets, please let us know! :)
Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
Prepare the sample DNA.
Place a 96-well skirted PCR plate on ice.
Add 10ng to 40ng of extracted DNA per well and bring to 30uL with nuclease-free water, or low TE.
Note
The 10ng to 40ng recommendation is a rough guideline. We have had successful experiments using less than 10ng; and also had reactions fail using 40ng. Probably more important than input DNA mass is sample quality and parasitemia.
Seal the plate and leave on ice while preparing the master mix.
Note
If you are using a cool block (e.g. Eppendorf 3881000031), be careful not to freeze the DNA while preparing the master mix.
Prepare the sWGA master mix.
| Reagent | Vol. per sample (uL) | Vol. per 96-well plate (uL) [+10% excess] | |
| Nuclease-Free H2O | 8.625 | 910.8 | |
| 10x Phi29 Buffer | 5 | 528 | |
| 20mg/mL BSA | 0.25 | 26.4 | |
| 1000uM sWGA Primer Mix | 0.125 | 13.2 | |
| 10mM dNTP | 5 | 528 | |
| phi29 (10U) | 1 | 105.6 | |
| TOTAL (uL) | 20 | 2112 |
Note
If working from a plate, aliquot master mix into a PCR strip tube and deliver by multichannel pipette to improve consistency.
Add 20uL of the master mix to each sample, mix by pipetting and incubate the plate in a thermal cycler with the following program:
| Step | Temp. (*C) | Time | |
| Amplify | 35 | 5 min | |
| 34 | 10 min | ||
| 33 | 15 min | ||
| 32 | 20 min | ||
| 31 | 30 min | ||
| 30 | 16 hr | ||
| Heat inactivation | 65 | 15 min | |
| Hold | 10 | Forever |
Note
The total reaction time is 17h35min. If you set the reaction up late afternoon (e.g. 3-4pm), it will be done for 9am the following morning.
(Optional) QC the sWGA product using a Qubit DNA assay. Good reaction performance will yield >20ng/uL; if your product is only at ~5ng/uL, it probably failed.
NOMADS Multiplex PCR
NOMADS Multiplex PCR
Transfer 2uL of sWGA product into a new skirted 96-well PCR plate.
Prepare KAPA HiFi polymerase (KK2101) master mix:
| Reagent | Vol. per sample (uL) | Vol. per 96-well plate (uL) [+10% excess] | |
| 5X Buffer with Mg2+ | 5 | 528 | |
| 10mM dNTPs | 0.75 | 79.2 | |
| Multiplex Primer Pool (10uM)* | 1.5 | 158.4 | |
| Nuclease-Free H20 | 15.25 | 1610.4 | |
| KAPA Pol. (1U/uL) | 0.5 | 52.8 | |
| TOTAL | 23 | 2428 |
*either NOMADS8 or NOMADS16 primers, as prepared in Section 1.
Note
If working from a plate, aliquot master mix into a PCR strip tube and deliver by multichannel pipette to improve consistency.
Note
We have also successfully amplified the NOMADS8 panel using the KAPA HiFi HotStart ReadyMix (KK2602). We haven't tried yet with the NOMADS16 panel; if you do and it works, let us know.
Mix by pipetting; or flick mix and spin down.
Add 23uL of master mix to each sample, and incubate the plate in a thermal cycler with the following program:
| Step | Temp. (*C) | Time | No. cycles | |
| Initial Denaturation | 95 | 3 mins | 1 | |
| Denaturation | 98 | 20 secs | 30 | |
| Annealing | 50 | 15 secs | ||
| Extension | 60 | 6 mins | ||
| Final Extension | 60 | 10 mins | 1 | |
| Hold | 8 | Forever | 1 |
Note
Why such a long extension time? According to the technical data sheet, KAPA HiFi extends at 15-60s/kbp. Our amplicons are 3-4kbp, so if we followed these guidelines a 4min extension should be adequate.
However, inspired by Xin-zhuan et al. (1996). NAR., we run with a reduced extension temperature (60*C rather than 72*C), which will decrease the rate of catalysis. With 6min we are trying to take this into account, and ensuring our longer and more difficult amplicons (e.g. crt1) are amplified.
Note
This is a safe stopping point. Store the PCR products at -20*C or 4*C.
(Optional) Run 2uL of PCR product on a 0.66% agarose gel. If you see a strong bands in the 3-4kbp range, this is a very good predictor that sequencing will be successful. If there are no bands visible in the 3-4kbp range, the multiplex PCR or sWGA has failed, and it is worth troubleshooting rather than continuing with sequencing.
SPRI bead clean-up of multiplex PCR products
SPRI bead clean-up of multiplex PCR products
Perform a 0.5X ratio SPRI bead clean-up of your multiplex PCR products.
Note
Ensure your SPRI beads are at room temperature!
Vortex your SPRI beads for 30s or more to ensure they are in suspension before pipetting!
Prepare a fresh stock of 80% ethanol each time.
To the PCR product, add 12uL of SPRI beads.
Mix thoroughly by pipetting or vortexing. Visually check to ensure the beads are mixed throughout the solution. If vortexing, spin down but do not pellet the beads.
Note
Especially when working in a 96-well plate, it is easy to inadvertently pipette the SPRI beads to the bottom of each well, pipette mix them at the bottom of the well, and therefore never mix them properly with the PCR product.
Incubate for 5mins at room temperature (RT).
Incubate on the magnet for 8mins.
Remove the supernatant. Do not transfer any beads; if this is a risk leave ~5uL of supernatant behind with the beads.
Wash the beads in 175uL of 80% ethanol. Leave for 30secs.
Remove and discard the supernatant. Repeat ethanol wash.
Spin down the samples, place back on the magnet and remove ay residual ethanol. Ensure all ethanol is removed.
Air dry the beads until the look dry, e.g. ~30secs.
Resuspend the beads in 15uL of nuclease-free water by pipetting.
Wait 2 to 5mins while DNA is being released from the beads. Waiting longer helps recovered longer DNA fragments.
Return samples to the magnet for 2mins or until beads have pelleted and the solution is clear.
Transfer 14uL of supernatant to a clean 96-well skirted plate.
QC using Qubit DNA assay. A good range is between 30-100ng/uL.
Note
We have observed a positive correlation between samples' ng/uL in this QC step and their P.falciparum mapping percentages in the sequencing data.
Note
This is a safe stopping point. Store at -20*C or 4*C.
One-pot barcoding by ligation (modified from Josh Quick's one-pot, see description)
One-pot barcoding by ligation (modified from Josh Quick's one-pot, see description)
Prepare the one-pot program on your thermal cycler, and start the program to bring the block to 20*C.
| Step | Temp. (*C) | Time | |
| Prepare block | 20 | Forever | |
| End prep. | 20 | 15 min | |
| Heat inactivate | 65 | 15 min | |
| Hold | 8 | Forever | |
| Prepare block | 20 | Forever | |
| Barcode ligation | 20 | 20 min | |
| Heat inactivate | 65 or 70 | 10 min | |
| Hold | 8 | Forever |
Note
If possible it is better to have the heated lid off for all steps that are not heat inactivation. If this is not possible on your thermal cycler, set the lid temperature lower (e.g 75*C) instead.
Transfer between 100-600ng of each sample's PCR product into a new plate, in no more than 10uL volume.
Note
You can attempt some sort of sample normalisation at this point, however in our experience ng of PCR product into the barcoding step only weakly influences final number of reads observed; so at present we feel it is preferable to transfer a consistent volume across the plate to minimise time and risk of cross-contamination.
Perform end preparation.
Prepare the end preparation master mix as follows:
| Reagent | Vol. (uL) per sample | Vol. (uL) per 96-well plate [includes 20% excess] | |
| NEBNext 10X End Prep Buffer | 1.4 | 161.3 | |
| NEBNext End Prep Enzyme Mix | 0.6 | 69.1 |
Add 2uL of the master mix to each sample. Mix by pipetting or flicking, and spin down.
Place the plate in the thermal cycler and press "Enter" to move the program forward; through the 20°C for 15 mins -> 65°C for 15 mins -> 8°C forever, steps.
Perform barcode ligation.
While the end preparation is incubating, thaw the native barcodes in the fridge. There may be condensation on the side of the barcode tubes requiring they are spun down.
Also while incubating, prepare the barcode ligation master mix:
| Reagent | Vol. (uL) per sample | Vol. (uL) per 96-well plate [includes 20% excess] | |
| Ultra II Ligase Master Mix | 6 | 691.2 | |
| Ligation Enhancer | 0.2 | 23 |
Once the thermal cycler block reaches 8*C, take out the plate and press "Enter" to move the program to the next 20*C hold.
Place the plate on ice for 1 min.
Note
The volume here is very small, if you are using a cool block (e.g. Eppendorf 3881000031), be careful not to freeze.
To each sample, add 0.5uL of the assigned native barcode.
Note
This step requires attention; double check your tips to ensure that 0.5uL of barcode is being delivered to every sample.
If you cannot consistently aspirate 0.5uL with your pipettes, you can transfer a larger volume (e.g. 0.75uL), although this increases assay costs.
Consider using long-reach thin-tipped pipette tips for this step. Inconsistencies in pipetting here directly influence sample balance!
Add 6.2uL of the barcode ligation master mix to each sample, mix well by pipetting and spin down.
Return the plate to the thermal cycler and press “Enter” to move the program through: 20°C for 20 mins -> 65°C for 10mins -> 8°C forever.
Sample pooling and SPRI bead clean-up.
Sample pooling and SPRI bead clean-up.
Pool equal volumes of each sample to a LoBind 1.5mL Eppendorf Tube.
Perform a 0.4X ratio SPRI bead clean-up.
Note
Ensure your SPRI beads are at room temperature!
Vortex your SPRI beads for 30s or more to ensure they are in suspension before pipetting!
Prepare a fresh stock of 80% ethanol each time.
To the pooled DNA, add SPRI beads at a 0.4X ratio.
Note
For example, if your pool is 100uL add 40uL of SPRI beads.
Mix thoroughly by pipetting or vortexing. Ensure the beads are mixed visually. If vortexing, spin down but do not pellet beads.
Incubate for 5 mins at RT.
Incubate on the magnet for 8 mins.
Remove the supernatant. Do not transfer any beads, if this is a risk leave ~5uL of supernatant behnid with the beads.
Wash the beads with 80% ethanol. Leave for 30 secs. Ensure the ethanol complete covers the pelleted beads – use an amount greater than the amount of sample + beads.
Remove and discard supernatant. Repeat ethanol wash.
Spin down the samples, place back on the magent and remove any residual ethanol. Ensure all ethanol is removed.
Air dry the beads until they look dry, e.g. ~30s.
Resuspend the beads in 70uL nuclease-free water by pipetting.
Wait 2 to 5 mins while DNA is released from the beads. Wait longer for longer DNA fragments.
Return samples to the magnet for 2 mins or until beads have pelleted and the solution is clear.
Transfer 67uL of supernatant to a clean plate.
QC 2uL of the cleaned pool using the Qubit.
Aim to pass 1 to 2ug (1000 to 2000ng) of pooled DNA into the adapter ligation reaction, making up to 65uL in nuclease-free water.
Library preparation and sequencing
Library preparation and sequencing
From this point forward, follow the protocol from Oxford Nanopore Technology (ONT)...
"1D Native barcoding genomic DNA (with EXP-NBD104, EXP-NBD114, and SQK-LSK109)"
...starting from the ‘Adapter ligation and clean-up’ step.
Make the following changes:
- Use LFB for the clean up
- Transfer 200-300fmol (~400-600ng) of DNA to the Flow Cell.