TITLE: Cost-effective targeted nanopore sequencing of P. falciparum malaria
AUTHORS: Mariateresa de Cesare, Mulenga Mwenda, Anna E. Jeffreys, Daniel J Bridges, Jason A Hendry

[DESCRIPTION]
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.

[STEPS]
SECTION: Preparation of primer pools.
1. Prepare the sWGA primer pool (see Materials).
SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
3. Prepare the sample DNA.


SECTION: Preparation of primer pools.
1.1. If you have ordered the primers lyophilised, make them up to 1000uM in nuclease-free water.
SECTION: Preparation of primer pools.
1.2. To make the 1000uM sWGA primer pool, combine equal volumes of each primer (e.g. Pf1-Pf10) in a 1.5mL LoBind tube.
SECTION: Preparation of primer pools.
1.3. Store the primers at 4C.
SECTION: Preparation of primer pools.
2. Prepare the NOMADS8 or NOMADS16 primer pools (see Materials).
SECTION: Preparation of primer pools.
2.1. If you have ordered the primers lyophilised, make them up to 100uM in nuclease-free water.
SECTION: Preparation of primer pools.
2.2. 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.

 

 

 

SECTION: Preparation of primer pools.
2.3. 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.

 

 



SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
3.1. Place a 96-well skirted PCR plate on ice.
SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
3.2. Add 10ng to 40ng of extracted DNA per well and bring to 30uL with nuclease-free water, or low TE.
 

SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
3.3. Seal the plate and leave on ice while preparing the master mix.

 

SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
4. Prepare the sWGA master mix.

 

 


SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
5. Add 20uL of the master mix to each sample, mix by pipetting and incubate the plate in a thermal cycler with the following program:
 

 


SECTION: Reduced cost sWGA (modified from Samuel O. Oyola et al. (2016))
5.1. (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.
SECTION: NOMADS Multiplex PCR
6. Transfer 2uL of sWGA product into a new skirted 96-well PCR plate.

SECTION: NOMADS Multiplex PCR
7. Prepare KAPA HiFi polymerase (KK2101) master mix:

 

 

 

SECTION: NOMADS Multiplex PCR
7.1. Mix by pipetting; or flick mix and spin down.
SECTION: NOMADS Multiplex PCR
8. Add 23uL of master mix to each sample, and incubate the plate in a thermal cycler with the following program:


 

 

 


SECTION: NOMADS Multiplex PCR
9. (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.
SECTION: SPRI bead clean-up of multiplex PCR products
10. Perform a 0.5X ratio SPRI bead clean-up of your multiplex PCR products.
 

SECTION: SPRI bead clean-up of multiplex PCR products
10.1. To the PCR product, add 12uL of SPRI beads.
SECTION: SPRI bead clean-up of multiplex PCR products
10.2. 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.
 

SECTION: SPRI bead clean-up of multiplex PCR products
10.3. Incubate for 5mins at room temperature (RT).
SECTION: SPRI bead clean-up of multiplex PCR products
10.4. Incubate on the magnet for 8mins.
SECTION: SPRI bead clean-up of multiplex PCR products
10.5. Remove the supernatant. Do not transfer any beads; if this is a risk leave ~5uL of supernatant behind with the beads.
SECTION: SPRI bead clean-up of multiplex PCR products
10.6. Wash the beads in 175uL of 80% ethanol. Leave for 30secs.
SECTION: SPRI bead clean-up of multiplex PCR products
10.7. Remove and discard the supernatant. Repeat ethanol wash.
SECTION: SPRI bead clean-up of multiplex PCR products
10.8. Spin down the samples, place back on the magnet and remove ay residual ethanol. Ensure all ethanol is removed.
SECTION: SPRI bead clean-up of multiplex PCR products
10.9. Air dry the beads until the look dry, e.g. ~30secs.
SECTION: SPRI bead clean-up of multiplex PCR products
10.10. Resuspend the beads in 15uL of nuclease-free water by pipetting.
SECTION: SPRI bead clean-up of multiplex PCR products
10.11. Wait 2 to 5mins while DNA is being released from the beads. Waiting longer helps recovered longer DNA fragments.
SECTION: SPRI bead clean-up of multiplex PCR products
10.12. Return samples to the magnet for 2mins or until beads have pelleted and the solution is clear.
SECTION: SPRI bead clean-up of multiplex PCR products
10.13. Transfer 14uL of supernatant to a clean 96-well skirted plate.
SECTION: SPRI bead clean-up of multiplex PCR products
11. QC using Qubit DNA assay. A good range is between 30-100ng/uL.
 

 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
13. Transfer between 100-600ng of each sample's PCR product into a new plate, in no more than 10uL volume.
 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
12. Prepare the one-pot program on your thermal cycler, and start the program to bring the block to 20*C.
 

 


SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
14. Perform end preparation.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
14.1. Prepare the end preparation master mix as follows:
 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
14.2. Add 2uL of the master mix to each sample. Mix by pipetting or flicking, and spin down.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
14.3. 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.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15. Perform barcode ligation.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.1. 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.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.2. Also while incubating, prepare the barcode ligation master mix:
 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.3. 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.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.4. Place the plate on ice for 1 min.
 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.5. To each sample, add 0.5uL of the assigned native barcode.
 

SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.6. Add 6.2uL of the barcode ligation master mix to each sample, mix well by pipetting and spin down.
SECTION: One-pot barcoding by ligation (modified from Josh Quick&#x27;s one-pot, see description)
15.7. 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.
SECTION: Sample pooling and SPRI bead clean-up.
16. Pool equal volumes of each sample to a LoBind 1.5mL Eppendorf Tube.
SECTION: Sample pooling and SPRI bead clean-up.
17. Perform a 0.4X ratio SPRI bead clean-up.
 

SECTION: Sample pooling and SPRI bead clean-up.
17.1. To the pooled DNA, add SPRI beads at a 0.4X ratio.
 

SECTION: Sample pooling and SPRI bead clean-up.
17.2. Mix thoroughly by pipetting or vortexing. Ensure the beads are mixed visually. If vortexing, spin down but do not pellet beads.
SECTION: Sample pooling and SPRI bead clean-up.
17.3. Incubate for 5 mins at RT.
SECTION: Sample pooling and SPRI bead clean-up.
17.4. Incubate on the magnet for 8 mins.
SECTION: Sample pooling and SPRI bead clean-up.
17.5. Remove the supernatant. Do not transfer any beads, if this is a risk leave ~5uL of supernatant behnid with the beads.
SECTION: Sample pooling and SPRI bead clean-up.
17.6. 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.
SECTION: Sample pooling and SPRI bead clean-up.
17.7. Remove and discard supernatant. Repeat ethanol wash.
SECTION: Sample pooling and SPRI bead clean-up.
17.8. Spin down the samples, place back on the magent and remove any residual ethanol. Ensure all ethanol is removed.
SECTION: Sample pooling and SPRI bead clean-up.
17.9. Air dry the beads until they look dry, e.g. ~30s.
SECTION: Sample pooling and SPRI bead clean-up.
17.10. Resuspend the beads in 70uL nuclease-free water by pipetting.
SECTION: Sample pooling and SPRI bead clean-up.
17.11. Wait 2 to 5 mins while DNA is released from the beads. Wait longer for longer DNA fragments.
SECTION: Sample pooling and SPRI bead clean-up.
17.12. Return samples to the magnet for 2 mins or until beads have pelleted and the solution is clear.
SECTION: Sample pooling and SPRI bead clean-up.
17.13. Transfer 67uL of supernatant to a clean plate.
SECTION: Sample pooling and SPRI bead clean-up.
18. QC  2uL of the cleaned pool using the Qubit.
SECTION: Sample pooling and SPRI bead clean-up.
18.1. 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.
SECTION: Library preparation and sequencing
19. 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.