May 15, 2026

Schistosoma haematobium larva FTA-to-WGS

DOI
https://dx.doi.org/10.17504/protocols.io.e6nvwwrp2vmk/v1
  • 1Natural History Museum;
  • 2Texas Biomedical Research Institute
  • pennance
Tom Pennance
Icon indicating open access to content
QR code linking to this content
DOI: https://dx.doi.org/10.17504/protocols.io.e6nvwwrp2vmk/v1
Protocol Citation: Tom Pennance, Daniel AJ Parsons, John Archer, Frédéric D Chevalier, Winka Le Clec'h, Timothy JC Anderson, Bonnie L Webster, Aidan M Emery 2026. Schistosoma haematobium larva FTA-to-WGS. protocols.io https://dx.doi.org/10.17504/protocols.io.e6nvwwrp2vmk/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: April 24, 2026
Last Modified: May 15, 2026
Protocol  Integer ID: 315662
Keywords: parasite, schistosome, schistosoma, urogenital schitosomiasis, library preparation, whole genome amplification, schistosoma haematobium larva fta, isolation of individual schistosome larva, individual schistosome larva, schistosoma haematobium, quantitative assessment of target parasite dna, target parasite dna, positive for schistosoma haematobium, collected larva, isolated cercaria from infected snail vector, larva, compatible whole genome sequencing, infected snail vector, preservation of larva, isolated cercaria, called miracidium, facilitating population genomic, urine egg microscopy, population genomic
Funders Acknowledgements:
Merck KGaA, Darmstadt, Germany
Grant ID: 10.13039/100009945
Disclaimer
DISCLAIMER – FOR INFORMATIONAL PURPOSES ONLY; USE AT YOUR OWN RISK

The protocol content here is for informational purposes only and does not constitute legal, medical, clinical, or safety advice, or otherwise; content added to protocols.io is not peer reviewed and may not have undergone a formal approval of any kind. Information presented in this protocol should not substitute for independent professional judgment, advice, diagnosis, or treatment. Any action you take or refrain from taking using or relying upon the information presented here is strictly at your own risk. You agree that neither the Company nor any of the authors, contributors, administrators, or anyone else associated with protocols.io, can be held responsible for your use of the information contained in or linked to this protocol or any of our Sites/Apps and Services.
Abstract
This protocol describes the isolation of individual schistosome larva, called miracidium, from urine samples determined positive for Schistosoma haematobium via urine egg microscopy (although this protocol could be followed for isolated cercaria from infected snail vectors), the preservation of larva on FTA cards, whole genome amplification (WGA) and quantitative assessment of target parasite DNA, and finally, the preparation of indexed Illumina-compatible whole genome sequencing (WGS) libraries.
The workflow enables the generation of high-quality sequencing-ready libraries from single field-collected larva, facilitating population genomic, epidemiological and drug resistance surveillance studies.
Materials
Consumables
Sample collection and egg hatching
  • 100 mL sterile urine collection pots
  • (optional for label printing) Zebra 2300 Performance Wax Ribbon 110mm x 74m
  • (optional for label printing) Zebra Z-Select 2000T Perfoation Paper Label 31x22mm
Urine egg microscopy
  • Plastic syringes (10–50 mL)
  • SEFAR NITEX 03-25/19 20 micron 13 mm diameter cold punched filter discs, (SEFAR cat. no. G060-1021-001-00)
  • Swinnex filter holder 13 mm (Merck cat no. SX0001300)
  • Glass microscope slide
  • Hydrophilic cellophane (soaked in glycerol)
  • 10% Lugol’s iodine solution
Schistosome egg hatching and miracidia collection
  • Plastic syringes, 20 micron 13 mm filter disc and filter holder as detailed under ‘Urine egg microscopy’
  • 9cm diameter Petri dishes
  • Fine forceps/tweezers
  • Bottled water or deionised water
  • QIAcard FTA Indicating Classic (100), 4 sample areas per card (Qiagen cat no. WB120206) (formerly Whatmann FTA indicating cards)
  • Sealable plastic bags for FTA storage
  • 20 / 200 µl micropipette tips
FTA punching and washing
  • Cutting mat for FTA cards (Qiagen cat no. WB100088/WB100020)
  • UniCore Punch 2.0 mm (Qiagen cat. no. WB100076) (formerly 2 mm Harris Micro-punch)
  • 1.5 mL microcentrifuge tubes (e.g. StarLab cat. no. S1615-5500 or other, such as 1.5 ml DNA LoBind tubes)
  • 0.2 mL PCR tubes or 8-strip PCR tubes (e.g. StarLab cat. no. A1402-3700)
  • 200 µl filtered pipette tips (e.g. StarLab S1120-8710)
  • 200 µl pipette tips (e.g. StarLab cat no. S1113-1700)
  • QIAcard FTA Wash Buffer (Qiagen cat no. WB120204 / WB120112)
  • TE-1 buffer (10 mM Tris-HCl, 0.1 mM EDTA), to make 100 ml, add:
  1. 1 mL 1M Tris-HCl, pH 8.0 (e.g. Invitrogen UltraPure 1M Tris-HCl, pH 8.0. ThermoScientific cat no. 15568025
  2. 20 µL 0.5M EDTA, pH 8.0 (e.g. Invitrogen UltraPure 0.5M EDTA, pH 8.0, ThermoScientific cat no. 15575020)
  3. 98.98 mL molecular grade water
Whole genome amplification (WGA)
  • 0.2 mL PCR tubes or 8-strip PCR tubes (e.g. StarLab cat. no. A1402-3700)
  • Cytiva illustra Ready-To-Go GenomiPhi V3 DNA Amplification Kit (Cytiva cat no. 25-6601-24 / 25-6601-96 / 25-6601-480)
  • Molecular biology grade water
  • 20/200 µl pipette tips (e.g. StarLab S1113-1700)
WGA magnetic bead purifications
  • 0.2 mL PCR tubes or 8-strip PCR tubes (e.g. StarLab cat. no. A1402-3700)
  • Beckman Coulter Agencourt AMPure XP magnetic beads (cat. no. A63880 / A63881 / A63882)
  • Freshly prepared 70% ethanol (molecular grade ethanol and water)
  • 25 ml reagent reservoirs (ThermoScientific cat. no. 8093)
  • 20/200 µl pipette tips (e.g. StarLab S1113-1700)
  • 10 µl pipette tips (e.g. StarLab S1111-3700)
qPCR and DNA quantification
  • 0.2 mL PCR tubes or 8-strip PCR tubes (e.g. StarLab cat. no. A1402-3700)
  • Qubit dsDNA BR Quantification Assay Kit (ThermoScientific cat no. Q32850 / Q32853)
  • 0.5 ml thin walled Qubit assay tubes (e.g. StarLab cat. no. I1405-8100)
  • Applied Biosystems SYBR Green Universal Master Mix (ThermoScientific cat. no. 4344463 / 4309155)
  • α-tubulin 2 forward primer (5′-GGTGGTACTGGTTCTGGTTT-3′)
  • α-tubulin 2 reverse primer (5′-AAAGCACAATCCGAATGTTCTAA-3′)
  • Cytiva PuReTaq Ready-To-Go PCR beads (Cytiva cat. no. 27955901)
  • Applied Biosystems MicroAmp Fast 8-Tube strip, 0.1 ml (ThermoScientific cat no. 4358293) and Applied Biosystems MicroAmp Optical 8-Cap Strips (ThermoScientific cat. no. 4323032) OR Applied Biosystems MicroAmp Optical 96-well Reaction Plate, (ThermoScientific cat no. 4316813) and Applied Biosystems MicroAmp Optical Adhesive Film (ThermoScientific cat no. 4311971)
  • 1000 µl pipette tips (e.g. StarLab S1122-1730)
  • 20/200 µl pipette tips (e.g. StarLab S1113-1700)
  • 10 µl pipette tips (e.g. StarLab S1111-3700)
  • (optional for DNA for standard curve generation) Qiagen Blood & Tissue kit
Library preparation
  • 0.2 mL PCR tubes or 8-strip PCR tubes (e.g. StarLab cat. no. A1402-3700)
  • New England Biolabs NEBNext Ultra II FS DNA Library Prep Kit for Illumina (NEB cat. no. E7805S / E7805L)
  • NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets 1–5 (NEB #E6440S/L, #E6442S/L, #E6444S/L, #E6446S/L, #E6448S/L)
  • 0.1X TE buffer (1X TE buffer comes with NEBNext library prep kit, dilute this 1:10 with water)
  • Freshly prepared 80% ethanol (molecular grade ethanol and water)
Library QC
  • Agilent D1000 ScreenTape Assay reagents, ladder and tips for TapeStation Systems (Agilent 5067-5582, 5067-5583, 5067-5599)
  • Qubit dsDNA BR Quantification Assay Kit (ThermoScientific cat no. Q32850 / Q32853)
  • (optional) KAPA Library Quantification Kits (Roche)

Equipment

Field equipment
  • Compound light microscope (for urine egg microscopy)
  • Dissection microscope (for capturing/isolating individual schistosome larva)
  • (optional for sample labelling) Zebra label printer – multiple machines available to suit 2000T label size.
  • Adjustable P20 micropipette

Molecular laboratory equipment
  • Adjustable micropipettes (P10, P20, P100, P200/300)
  • (optional) Adjustable multichannel micropipettes (P10, P20, P100, P200/300)
  • 1.5 ml tube holder/rack
  • 0.2 ml tube holder
  • Vortex mixer
  • Mini centrifuge (e.g. Starlab Minicentrifuge cat no. S8030-5000)
  • Plate centrifuge
  • Tube rocker or rocking platform
  • Dry bath incubator / heat block capable of 50°C (e.g. LabNet AccuBlock)
  • Wet ice bucket / wet ice machine
  • Magnetic rack for 0.2 mL tubes (e.g. Sergi Lab Supplies magnetic rack for DNA, RNA purification, cat. no. 1018b)
  • (optional) Microbead sterilizer (e.g. Steri 250, Sigma-Aldrich cat. no. Z378569)
Thermal cycling and amplification equipment
  • Thermal cycler with adjustable heated lid (e.g. Bio-Rad C1000 Touch Thermal Cycler)
  • qPCR instrument (e.g. Applied Biosystems QuantStudio)
DNA quantification and library QC
  • Qubit Fluorometer
  • Agilent TapeStation
Schistosoma haematobium diagnostic
Urine sample from patient collected in an individual sample pots (50 ml) labelled with the participant ID code. Fresh urine samples are kept in a cooler (or fridge) and in the dark until processing
Urine filtration microscopy for S. haematobium egg detection performed as described in Knopp et al. BMC Infectious Diseases (2026)26:263.
Schistosoma haematobium egg hatching and miracidia collection
12h 48m
Immediately prior to use, the positive urine sample is homogenised by vigorous mixing or using the 10ml urine filtration syringe (urine is repeatedly drawn in, and then pushed, out of the syringe).
1m
All of the remaining homogenised urine is taken up into the syringe (in stages depending on volume of syringe and remaining urine volume, until all urine is used). The urine is then pushed through the SEFAR Nitex 03-25/19, 20 micron 13 mm stamped disk (filter) housed inside the Swinnex filter holder (13 mm) with any S. haematobium eggs (size range 112-170 µm x 40-70 µm) being captured on the filter.
1m
The filter holder is carefully opened and the filter is carefully removed using a clean pair of forceps and placed in a clean, labelled, petri dish containing clean bottled or deionised water.
1m
The lid of the petri dish is put on and then the petri dish is put in a warm environment in indirect sunlight (so not to get too hot) or under a lamp to stimulate schistosome egg hatching and miracidia swimming.
Note: Some samples will start to hatch quite quickly, but slower ones may take ~2 hours, so keep checking plates for presence of motile miracidia using a dissection microscope.
9cm Petri dishes containing urine filters and bottled/deionised water placed in indirect sunlight to stimulate egg hatching.

30m
Using a dissection microscope, schistosome miracidia are then individually captured in 3µl using a p20 pipette. The 3 µl of liquid containing the miracidium is then pipetted directly on to a labelled FTA card by gently placing the tip on the card surface (to avoid tearing or pressing the card material) and expelling the liquid onto the card surface, ensuring enough space is left between each miraicida 'spot'. See video and the pictured FTA card in the next step for reference.

Video
Video demonstrating the capturing of individual schistosome miracidia using a dissection microscope and p20 micropipette, and placement of miracidium on an FTA card for DNA capture.
Note: This video shows capturing S. mansoni miracidia from a stool sample, for S. haematobium there will only be a 20 micron 13 mm urine filter in the petri dish and not stool carryover from a pitchford funnel.
Video credit: Aidan M. Emery, Schistosomiasis Collection at the Natural History Museum.
15m
Allow the FTA cards to dry overnight (or for a few hours at least) away from direct sunlight by propping the pink coloured capture material side of the FTA up on its cover (see picture). Once dried, fold over the protective cover and seal FTA cards together in a sealable plastic bag to protect from moisture, and store in a cool, dark place, in another sealed container.
FTA cards with captured schistosome miracidia in a drying position

12h
Whole genome amplification (WGA) of Schistosoma haematobium DNA stored on FTA
3h
Placing the fully unfolded FTA card on a cutting mat, punch a 2 mm spot containing miracidium using a UniCore Punch 2.0 mm and transfer to a labelled 1.5mL microcentrifuge tube.
FTA card containing schistosome miracidia, some of which have been punched out and placed into 1.5 ml tubes.

5m
Using a p200 pipette and filtered 200 µl pipette tip, add 180 µl of QIAcard FTA wash buffer, vortex very briefly to mix, and incubate for 5 minutes at room temperature on a mixer (low rpm) or rocking platform.
Rocking platform with 1.5 ml tubes in a rack, each containing an FTA punch and purification reagent. Note: Tube lids are open merely to save time between capping/uncapping.
Note 2: Tube lids can also remain open during the brief vortex of the 180 µl + FTA punch if performed correctly. This takes some practice, but uses the fluid dynamics to its advantage.

5m
Using a p200 pipette (set to 200) and unfiltered 200 µl pipette tip, remove and discard FTA purification reagent and tip (minimising contact with punch).
1m
Repeat the FTA purification reagent wash and removal step.
Note: Increase to repeating twice/three times depending on sample type and quality (i.e. if substandard results downstream, this may suggest card contains excess inhibitors that could be removed using multiple washes).
5m
Using a p200 pipette and filtered 200 µl pipette tip, add 180 µl of TE-1 buffer, vortex very briefly to mix, and incubate for 5 minutes at RT on a mixer (low rpm) or rocking platform.
5m
Using a p200 pipette and unfiltered 200 µl pipette tip and 200 pipette set to 200, remove and discard TE-1 buffer and tip (minimising contact with punch).
1m
Repeat the TE-1 wash and removal step.
Note: Increase to repeating twice/three times depending on sample type and quality (i.e. if substandard results downstream, this may suggest card contains excess inhibitors that could be removed using multiple washes)
5m
On the second removal of TE-1, use the pipette tip to gently place FTA punch on the side of tube (see picture) to help separate the punch from any residual liquid at the bottom of the tube and aid in the following drying step.
FTA card punch placed on the side of 1.5 ml tube to move away from any residual liquid at the bottom of the tube.

1m
Dry punches until damp (do not overdry) in the microcentrifuge tubes with lids open at 50C for 5-10 minutes in a dry bath / heat block.
Dry bath incubator set to 50ºC with two tubes containing an FTA punch.

5m
Once the FTA punch is just slightly damp carefully transfer the punch to a 0.2 mL PCR tube, using a pair of fine tweezers.
Note: clean the tweezers between each sample transfer using 70% ethanol and a lint-free tissue wipe, ensuring removal of any residual FTA card fibres or liquid. For ultra-conservative contamination control, additionally use a microbead heat sterilizer or a low percent (10%) bleach solution.
2m
Make the Cytiva denaturation buffer solution by pre-mixing (per sample):
  • 10 µl ultra pure water
  • 10 µl GenomiPhi denaturation buffer (provided in Cytiva GenomPhi V3 kit)
1m
Add 20 µl of the denaturation buffer solution to the 0.2ml tube containing the FTA punch. Briefly centrifuge down to ensure FTA punch is submerged in the denaturation buffer.
Heat the 0.2ml tube containing the FTA punch/denaturation buffer to 95ºC for 3 minutes (denatures DNA) by adding tubes directly to a pre-heated thermal cycler. Do not set any cooling step on the PCR machine, instead, after 3 minutes at 95ºC, move the tubes directly from the PCR machine and immediately onto wet ice (in a 0.2 ml tube holder, see picture in next step) to avoid DNA renaturation.
3m
While on ice, remove the 20 µl denatured buffer from FTA punch and add to the Cytiva Ready-To-Go GenomiPhi V3 WGA tube containing the enzymatic cake (see picture), seal the lids using the Cytiva supplied caps, and wait for enzymatic cake to dissolve (5-10 minutes) whilst doing intermittent finger flicks / gentle vortexing to mix sample and aid the dissolving of the enzymatic cake, keeping cool throughout.
During liquid handling phases, a p100 multichannel pipette with 200 µl unfiltered tips can be used to transfer the 20 µl denaturation buffer & denatured DNA away from the tubes containing the FTA card (left) to the Cytiva tubes (right) containing the enzymatic cake. The tubes containing the FTA card are retained on ice during this incubation step.
Note: The Cytiva GenomPhi V3 96 well plates can be seperated into columns along the perforated edges and carefully using a scalpel/blade to cut through the foil seal.

10m
Once the enzymatic cake is fully dissolved, spin down the dissolved enzymatic cake briefly (i.e. in a plate centrifuge for 2 seconds at 1,000 xg) and transfer the dissolved enzyme cake liquid (20 µl) back to the tube containing the FTA punch, spin down briefly, and incubate in PCR machine with the following steps:

  1. 30 ºC for 120 minutes
  2. 65ºC for 10 minutes
  3. 4ºC hold for ~infinity

Ideally, proceed immediately to the next step once the incubation is complete (i.e. at 4ºC hold step), but if necessary, the reaction can be left overnight at 4ºC hold in the machine without seemingly impeding downstream results.
2h 10m
After removing tubes from the thermal cycler, EITHER:

Transfer the 20 µl of WGA product liquid (i.e. leave behind FTA card punch that can be discarded with the reaction tube) to a new 0.2 ml PCR strip and store at -20C avoiding freeze-thaw cycles,

OR;

If proceeding directly to WGA purification, transfer the 20 µl of WGA product liquid (i.e. leave behind FTA card punch that can be discarded with the reaction tube) to a 0.2 ml PCR strip containing 36 µl of room temperature AMPure XP beads (see next section for details on preparing these tubes in advance). Discard the reaction tube containing the FTA punch.

1m
WGA product purification (magnetic beads)
52m
Remove Beckman Coulter Agencourt AMPure XP beads from fridge and warm to room temperature. Vigorously shake and vortex to resuspend the beads in solution and make homogenous.
30m
Pipette 36 µl (1.8 x of DNA sample volume) of beads into new 0.2ml PCR tubes.
1m
Transfer the 20 µl of WGA product liquid (i.e. leave behind FTA card punch that can be discarded with the reaction tube) from the WGA reaction tube to the new tube containing the magnetic beads.
1m
Briefly vortex (or pipette mix, set pipette to 30 µl and pipette ~10 times) WGA product and magnetic beads until homogenous, give a quick spin down, and incubate at room temp for 5 minutes.
Reminder: during this incubation period, prepare the 70% ethanol used in the proceeding wash steps (360 µl per sample).
5m
After 5 minutes, move tubes to a 0.2 ml magnetic rack for 2 minutes until solution clears.
2m
Set p100/p200 to >56 µl and remove and discard supernatant only (do not touch bead side of tube).
1m
Add 180 µl of freshly made 70% ethanol to tube containing beads and incubate for 30 seconds at room temperature. Ensure ethanol volume is enough to cover the beads for incubation time.
Note: Use reagent reservoirs for multichannel dispensing.
30s
Remove ethanol (set to 200 µl).
1m
Repeat 70% ethanol wash and removal steps once, for a total of two times. On final removal, use a p10 if necessary to remove any remaining ethanol from the bottom or sides of tubes to ensure as little ethanol carry over as possible.
Caution: Once ethanol removed, proceed immediately to next step and do not allow tubes to sit without ethanol for >2 minutes, which may cause the beads to overdry. Ensure beads retain a glossy finish, if they start to dull in colour and crack, the beads are too dry.
30s
Remove tube from magnetic rack, and add 31 µl Molecular Biology Grade water and gently vortex (or pipette mix, set pipette to 20 µl and pipette repeatedly the bead mixture onto magnetic bead pellet) until the bead pellet has disappeared from the side of tube and all beads are in solution.
Note: Resuspending the bead pellet in water may take longer if the beads are left to over dry following ethanol removal, so it is important to get the timing of this step right.
2m
Once beads are homogenous in the water (no remaining bead pellet on side of tube), incubate for 5 minutes at room temperature.
5m
After 5 minutes, move the tube to magnetic rack for 1 minute until solution clears (should be quite quick).
1m
Transfer 30 µl eluate to new tube without disturbing the pellet, or carrying over any pellet (caution should be taken as carry over may happen during the final few µl of pipetting.
1m
Make a 100 fold dilution of the purified WGA product if proceeding to Qubit and qPCR quantification steps. To do this add 198 µl of ultra pure water to a new 0.2 ml PCR tube and then add 2 µl of the purified WGA product.

1m
Store remaining ~28 µl of purified WGA product in the labelled 0.2 ml tubes at -20ºC until ready ready to proceed with library preparation steps.
Qubit quantification
20m
This protocol is as described by the invitrogen Qubit dsDNA BR Assay Kit User Guide:
  1. Prepare the Qubit working solution by diluting the Qubit dsDNA BR Reagent 1:200 in Qubit dsDNA BR Buffer (e.g. for 10 samples, add 10 µl of Qubit dsDNA BR Reagent to 1,990 µl of Qubit dsDNA BR Buffer and mix.
  2. Add 190 µl of Qubit working solution to each 0.5 ml assay tube (StarLab 0.5 ml PCR Tube, Flat Cap, Natural cat no. I1405-8100).
  3. Add 10 µl of the 100 fold diluted purified WGA product to each assay tube and vortex for 3-5 seconds.
  4. Read assay tube concentration on Qubit and back calculate WGA concentration by multiplying by 100.
20m
qPCR quantification of Schistosoma haematobium WGA
6h 26m
Generate standard curve and efficiency calculation (skip to the next step if complete)

Before conducting qPCR for quantification of whole genome copy numbers per S. haematobium sample, it is necessary to generate a standard curve to back calculate qPCR reaction cycle numbers with genome copy number. There are several protocols on how to conduct this. The following was expanded and adapted from work originally conducted by Winka Le Clec'h and Frédéric Chevalier (Texas Biomedical Research Institute) using the single-copy locus, S. haematobium α-tubulin 2 (Sh α2), as described briefly in:
Le Clec’h W, Chevalier FD, McDew-White M, et al. Whole genome amplification and exome sequencing of archived schistosome miracidia. Parasitology. 2018;145(13):1739-1747. doi:10.1017/S0031182018000811.
The starting material for the standard curve generation was a Schistosoma haematobium adult worm obtained from the Schistosomiasis Collection at the Natural History Museum, the DNA of which was extracted using the Qiagen Blood and Tissue kit following manufacturers instructions, and DNA eluted in 50 µl of ultra pure water.
25 µl of PCR product using the Sh α2 primers was generated using Cytiva PuReTaq Ready-To-Go PCR beads with the following reaction mixture:
  • 1 µl - Sh α2 forward primer 10 µM (5′-GGTGGTACTGGTTCTGGTTT-3′)
  • 1 µl - Sh α2 reverse primer 10 µM (5′-AAAGCACAATCCGAATGTTCTAA-3′)
  • 21 µl - molecular biology grade water
  • 2 µl - S. haematobium adult worm DNA

With the following cycling conditions:
ABC
TEMPERATURETIMECYCLES
95ºC5 minutes1
95ºC30 secondsx 35
60ºC30 seconds
72ºC30 seconds
72ºC10 minutes1
4ºCinfinityhold
PCR cycling conditions for S. haematobium α-tubulin 2 (Sh α2) reaction

1h 20m
The 25 µl PCR product is then bead cleaned by adding 45 µl (1.8x) of Beckman Coulter Agencourt AMPure XP beads (warmed to room temperature), following the same incubation and 70% ethanol wash steps as described for the WGA product purification (i.e. 2 x 180 µl 70% ethanol washes, see above). The final elution of purified PCR product was performed with 30 µl of pure water.
10m
Perform QuBit DS DNA Broad Range quantification using 2-5 µl of PCR product in three separate tubes (i.e. 3 replicates). Take the mean of these 3 measurements as the purified PCR product concentration to input into the yellow cell of the attached qPCR standards calculator provided by Winka Le Clec'h and Frédéric Chevalier.
Download qPCR_standards_Sh_alpha-tubulin.xlsxqPCR_standards_Sh_alpha-tubulin.xlsx11.7KB

10m
Generate the standard serial dilutions starting from standard S1 (diluting 1 part PCR product in 39 parts water). Recommend to use same volume as was used in the QuBit calculations for this S1 dilution (e.g. 2:78 µl or 5:195 µl). Then generate 10 fold dilutions to generate S2 - S7 (i.e. 1 part S1 to 9 parts water = S2, 1 part S2 to 9 parts water = S3, and so on...)
5m
Perform standard curve qPCR (follow qPCR reaction set up and cycle as described below) for the standards S1-S7 (run each standard in triplicate), and allow analysis program (e.g. applied biosystems Design & Analysis software 2.6.0) to automatically calculate a suitable fluorescence (∆Rn) to record the cycle threshold across standards. You will use this same threshold across all test samples to maintain consistency and comparison to standards (for example 0.726 as shown below).

qPCR amplification plot of standards S2-S6 showing ∆Rn value of 0.726 that the Applied Biosystems Design & Analysis software 2.6.0 predicts is in a consistent exponential amplification point for each reaction.

Also, ensure that melt curve is consistent across all reactions. Any non-uniform peaks in the melt curves indicate non-target amplification (perhaps due to contamination or insufficient bead cleaning etc) and, if present, should be repeated as this will affect downstream genome copy calculations

qPCR melt curve plot of standards S2-S6 showing showing a consistent melt temperature peak between ~74.0ºC

As above, removal of highest (S1) and lowest (S7) standards may be required to obtain an R2 = 1.000 (or as close to 1 as possible) and a qPCR efficiency (E) = 100%. Triplicate readings for each standard should be identical, but if an obvious outlier exists, remove the outlier, but ensure that at least two points remain per standard.

Once final standards to include in analysis are established, record the values for slope and Y-intersect from the standard curve plot, which will later be used to back-calculate the S. haematobium genome copy number for your test samples.

qPCR standard curve plot of standards S2-S6 triplicate readings, showing good fit of data (R2=0.996) and qPCR efficiency (101.797%).
Note: This protocol describes the generation of a single standard curve that is then used for all subsequent qPCR experiments. While this is not uncommon, other users may prefer to run standards and generate a standard curve on each plate of samples they run, an approach which although does take more time and consumes more reagents, does allow you to account for variations in the PCR mix across plates. The diluted standards S1-S7 can be kept for up to ~six months at -20ºC to be used across experiments.
2h
Sample qPCR and genome copy number calculation

Before setting up qPCR reaction, input the run method and plate set up (sample information) in the qPCR software and load onto the machine so that when the plate is ready, it can be loaded immediately into the machine. The Instrument used here is a QuantStudio 6 Pro with a 96-Well 0.1 mL Block.

Open qPCR software (e.g. Applied Biosystems Design & Analysis software 2.6.0), select:
Run Mode: Standard (i.e. not fast)
Analysis Module: Standard Curve

Proceed to Run Method, input:
Reaction volume = 10.0 µl
Heated cover temperature 105.0ºC
and the following cycling parameters:

qPCR run method for S. haematobium α-tubulin 2 (Sh α2)

Proceed to Plate Setup, input:
Samples - desired sample Name(s) (Type = Unknown) for those being run and include a negative control (Type = Negative Control) and allocate to respective wells
Targets - Set Target for all as 'a-tubulin' and the Reporter as 'SYBR'

Proceed to sending the Run to the instrument and 'Load Plate File' on the machine.

Note: Other machines may differ in their setup.

10m
Set up qPCR reaction

In a pre-PCR hood, set up a qPCR reaction master mix in an appropriate size tube, per reaction:
  • 5 µl Applied Biosystems SYBR Green Universal Master Mix (cat no. 4309155)
  • 3.4 µl Ultra pure water
  • 0.3 µl α-tubulin 2 forward primer
  • 0.3 µl α-tubulin 2 reverse primer

Vortex master mix and pipette 9 µl into each optical qPCR tube or well of optical qPCR plate.
10m
Add DNA input

For each WGA'd sample, use 1 µl of the 100 fold diluted purified WGA product* generated earlier and add to the qPCR reaction, seal the tube/plate using Applied Biosystem tube caps or plate MicroAmp Optical Adhesive Film and centrifuge down briefly (1 min at 2000 xg to remove and bubbles), proceed immediately to run the qPCR.

*Note: Once 100 fold dilution of WGA product is made, it can be stored for a few days at -20ºC and used, but following which it is recommended to generate a new dilution from the WGA product).
1m
Run the qPCR on an Applied Biosystems QuantStudio (or similar) with the aforementioned run method.
2h
Once complete, open the experiment results file and change the ∆Rn threshold for the target a-tubulin to your pre-determined ∆Rn value (e.g. 0.726). On the Applied Biosystems Design & Analysis software 2.6.0, this option can be found under Actions > Primary Analysis Settings. Deselect 'Use default' and 'Auto Threshold' and input your Threshold value, see screenshot:


Once done, click save and then 'Analyze', and wait for the program to complete the analysis using the new threshold value. Once finished, click Actions > Export, and save the results as your desired file format (e.g. a .xlsx in a 'Combined all export tables in one file')
10m
Analysis of qPCR data

Open the exported qPCR data from the experiment, select the 'Results' file/tab where you will find details of the quantification (Cq) and primary melt curve temp (Tm1) for each sample. Disregard samples with a Tm1 that falls outside of the expected range (e.g. <73.8>74.8)

For each sample, calculate the genome copy number for each reaction using the formula:

=10^((Cq - Y-intercept) / slope)

!!! NB: Y-intercept and slope were obtained from your standard curve generation and stay the same for all test samples. !!!

Since using a 100 fold dilution of the original sample WGA, multiply this value by 100 to get your WGA genome copy number.
10m
Determine S. haematobium copies per ng of DNA and sample selection for library preparation

A simple calculation:
WGA genome copy number (qPCR)
/
WGA concentration (QuBit)
=
S. haematobium genome copies per 1ng of DNA (Sh copies/ng)

The higher the Sh copies/ng, the better your target sequencing coverage should be (technically). Although any sample that has amplified the S. haematobium target in the qPCR can be selected, preferentially select samples with the highest Sh copies/ng to increase sequencing efficiency and target read depth.

N.B. Given that 1 picogram of DNA contains an estimated 978,000,000 base pairs, the S. haematobium genome (385 Mb) weighs 0.39 pg (haploid), therefore 1 ng of 100% S. haematobium genomes should have a maximum of 2,564 (1000/0.39) genome copies. However, WGA products can provide Sh copies/ng results far above this, which can be caused by WGA artifacts (qPCR target may be overrepresented), small Cq shifts on qPCR from pipetting errors or slight qPCR efficiency overestimation (E > 100%, see standard curve plot above) or other stochasticity. Remember, the purpose here is not to get a perfect estimation of genome copy number, but to try and select samples that will (hopefully) return the highest proportion of target DNA in libraries during the next step.
Library preparation for each selected whole genome amplified sample
2h 59m 30s
Input for library generation can vary, and for the New England Biolabs NEBNext Ultra II FS DNA Lib Prep Kit for Illumina used here, has protocols for use with inputs <100 ng and >100 ng, which should be easily achieved following WGA.

The following steps are adapted from the original NEBNext Ultra II FS DNA Lib Prep Kit for Illumina instruction manual version 4.0_7/23 (manualE6177-E7805.pdf), Section 2: Protocol for use with Inputs ≥100 ng, to generate libraries indexed with NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets 1–5 for pair ended 150 bp sequencing on an Illumina platform.

Note: Unless otherwise stated, thaw all NEBNext reagents on ice.
Based on the QuBit quantification result, generate the input DNA starting material of between 100-200 ng of the purified WGA product in a total volume of 26 µl, normalising the DNA amount in ultra pure water to achieve desired input. Typically, this input can be achieved by a 25, 50 or 100 fold dilution of the WGA product, or normalise each product to the same ng.
Fragmentation/End prep
Ensure that the Ultra II FS Reaction Buffer is completely thawed. If a precipitate is seen in the buffer, pipette up and down several times to break it up, and quickly vortex to mix. Place on ice until use.
1m
Vortex the Ultra II FS Enzyme Mix 5-8 seconds prior to use and place on ice.
Note: It is important to vortex the enzyme mix prior to use for optimal performance.
30s
Add the following components to a 0.2 ml thin wall PCR tube on ice:
AB
COMPONENT VOLUME PER ONE LIBRARY
DNA*26 µl
• (yellow) NEBNext Ultra II FS Reaction7 µl
• (yellow) NEBNext Ultra II FS Enzyme Mix 2 µl
Total Volume 35 µl
*Use 100-200 ng purified WGA product
Vortex the reaction for 5 seconds and briefly spin in a mini centrifuge.
30s
In a thermal cycler, with the heated lid set to 75ºC, run the following program:
  1. 20 minutes @ 37ºC (20 minute incubation aimed for fragment size of ~300-350 bp)
  2. 30 minutes @ 65ºC
  3. Hold @ 4ºC
50m
Recommendation is to continue directly to the next step (Adapter Ligation), though if necessary, samples can be stored at -20ºC overnight although a loss of yield may be observed (~20% reported by NEB)
Adapter Ligation

Note: Prior to starting, ensure that the following components are thawed on ice:
NEBNext Ultra II Ligation Master Mix
NEBNext Ligation Enhancer
NEBNext Adaptor for Illumina
NEB USER Enzyme (comes with the NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets)
Add the following components (premixing is possible, see Note below) directly to the fragmented DNA mixture:

AB
COMPONENT VOLUME
Fragmented DNA (from Fragmentation/End Prep step)35 µl
• (red) NEBNext Ultra II Ligation Master Mix* 30 µl
• (red) NEBNext Ligation Enhancer 1 µl
• (red) NEBNext Adaptor for Illumina** 2.5 µl
Total Volume 68.5 µl
* Mix the Ultra II Ligation Master Mix by pipetting up and down several times prior to adding to the reaction.
** The NEBNext adaptor is provided in NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets.
Note: The Ligation Master Mix and Ligation Enhancer CAN be mixed ahead of time and is stable for at least 8 hours @4°C. For example, for 8 library preps, mix 240 µl Ligation Master Mix and 8 µl Ligation Enhancer.
Do not, however, premix with the Adaptor for Illumina until directly prior to use in this Adaptor Ligation Step. For example, for 8 library preps, add the 20 µl to the 248 µl Ligation MM/Enhancer immediately before adding to fragmented DNA.
10m
Once added to the fragment DNA, set a 100 µl or 200 µl pipette to 50 µl and pipette the reaction mixture volume up and down at least 10 times to mix thoroughly. Perform a quick spin to collect all liquid from the sides of the tube.
Caution: The NEBNext Ultra II Ligation Master Mix is very viscous. Care should be taken to ensure adequate mixing of the ligation reaction, as incomplete mixing will result in reduced ligation efficiency. The presence of a small amount of bubbles will not interfere with performance.
1m
Incubate at 20°C for 15 minutes in a thermal cycler with the heated lid off.
Reminder: Remove AMPure XP magnetic beads from fridge and warm to room temperature (~30 minutes) for use in proceeding Size Selection.
15m
Once incubation complete, remove from thermal cycler and immediately add 3 µl of the NEB USER Enzyme to the ligation mixture
1m
Mix well (set a 100 µl or 200 µl pipette to 50 µl and pipette the reaction mixture volume up and down at least 10 times to mix thoroughly) and incubate at 37°C for 15 minutes with the heated lid set to 47°C.
15m
Recommendation is following incubation to proceed directly to size selection, but that adapter ligated DNA can be stored overnight at -20ºC.
Size Selection of Adaptor-ligated DNA for DNA Input ≥ 100 ng

Note: ensure that AMPure XP beads have warmed to room temperature before proceeding to size selection. Size selection (and therefore bead ratios) below aims to target final library sizes of 320-470 bp, (following the original insert size selection of 300-350 bp).
Bring the volume of the reaction up to 100 µl by adding 28.5 µl 0.1X TE (dilute 1X TE Buffer 1:10 with water).
1m
Vortex AMPure XP beads to resuspend.
1m
Add 30 μl (~ 0.3X) of resuspended beads to the 100 μl sample, vortex for 3-5 seconds on high. Centrifuge briefly after mixing (e.g. 1 second) to collect liquid/beads from sides of tube, but be sure to stop the centrifugation before the beads start to settle out.
1m
Incubate samples for at least 5 minutes at room temperature.
5m
Place the tube on 0.2 ml magnetic stand to separate the beads from the supernatant for 5 minutes.
5m
After 5 minutes (or when the solution is clear), carefully transfer the supernatant (~ 130 µl) containing your DNA to a new 0.2 ml thin wall PCR tube i.e. Caution: do not discard the supernatant! Discard the beads that contain the unwanted large fragments.
Add 15 μl (~0.15X) resuspended AMPure XP to the supernatant and vortex for 3-5 seconds on high. Centrifuge briefly after mixing (e.g. 1 second) to collect liquid/beads from sides of tube, but be sure to stop the centrifugation before the beads start to settle out.

Incubate samples for at least 5 minutes at room temperature.

Reminder: Remove the desired NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Set from the freezer and leave to thaw for 10 to 15 minutes
5m
Place the tube on an appropriate magnetic stand for 5 minutes to separate the beads from the supernatant.

Reminder: During this incubation step, prepare some fresh 80% ethanol for proceeding bead cleaning. You will need ~360-400 µl per library prep for cleaning up adapter ligated DNA, and you will ALSO need an additional ~360-400 µl per library prep for cleaning up the PCR enriched library later on in the protocol.
5m
After 5 minutes (or when the solution is clear), carefully remove and discard the supernatant that contains unwanted DNA. Caution: Be careful not to discard/disturb the beads that contain the desired DNA.
Add 180 μl of 80% freshly prepared ethanol to the tube/plate while in the magnetic stand. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant (setting pipette to 200 µl). Be careful not to disturb the beads that contain DNA targets.
1m
Repeat the 80% ethanol wash step once for a total of two washes. Be sure to remove all visible liquid after the second wash. If necessary, briefly spin the tube/plate, place back on the magnet and remove traces of ethanol with a p10 pipette tip.
1m
Air dry the beads for up to 5 minutes while the tube/plate is on the magnetic stand with the lid open.
Caution: Do not overdry the beads. This may result in lower recovery of DNA. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack, they are too dry.
5m
Remove the tube/plate from the magnetic stand. Elute the DNA target from the beads into 17 μl 0.1X TE (dilute 1X TE Buffer 1:10 in water).
2m
Mix well on a vortex mixer and incubate for at least 2 minutes at room temperature, ensure beads from side of tube are resuspended. If necessary, quickly spin the sample to collect the liquid from the sides of the tube or plate wells before placing back on the magnetic stand.
2m
Place the tube/plate on a magnetic stand. After 5 minutes (or when the solution is clear), transfer 15 μl to a new PCR tube.
Optional: instead of to a new tube, transfer the adapter ligated DNA directly to the PCR enrichment tube containing the PCR enrichment master mix/index primer mix as described in the 'PCR Enrichment of Adapter-ligated DNA' step if proceeding immediately.
5m
Proceed directly to PCR Enrichment of Adaptor-ligated DNA or samples can be safely stored at -20ºC.
PCR Enrichment of Adapter-ligated DNA

Note: Prior to starting, ensure that the following components are thawed:
(on ice) NEBNext Ultra II Q5 Master Mix
(room temp for 15 minutes) Index Primer Mix (NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Set)
Centrifuge the NEBNext Indexes plate (280 × g for 1 min) to collect all of the primer at the bottom of each well.
1m
To new 0.2 ml PCR tube strips add the 25 µl Q5 master mix and the 10 µl primer index (use tips to pierce the foil seal on the plate of indexes to keep track of those used), and then add the adaptor ligated DNA to each. The table summarises this reaction set up:
AB
COMPONENT VOLUME
Adaptor Ligated DNA Fragments (from previous step)15 µl
• (blue) NEBNext Ultra II Q5 Master Mix 25 µl
• (blue) Index Primer Mix* 10 µl
Total Volume 50 µl
*From the NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets

5m
Seal tubes and gently vortex or pipette mix the 50 µl reaction mixture thoroughly and quickly centrifuge to collect all liquid from the side of tubes.
1m
Place tubes in a thermal cycler with the lid set to 105ºC and perform PCR enrichment using the following conditions:
ABCD
CYCLE STEP TEMP TIME CYCLES
Initial Denaturation 98°C 30 seconds 1
Denaturation 98°C 10 seconds4
Annealing/Extension 65°C 75 seconds
Final Extension 65°C 5 minutes 1
Hold 4°C∞
Reminder: Remove AMPure XP beads for proceeding clean up from fridge and allow to warm to room temperature.
12m
Bead clean PCR enriched library
Vortex the room temperature AMPure XP beads to resuspend.
30s
Add 45 μl (0.9X) resuspended AMPure XP beads to the PCR reaction. Vortex for 3-5 seconds on high, centrifuge briefly (1 second) to collect the liquid from the sides of the tube, be sure to stop the centrifugation before the beads start to settle out.
1m
Incubate samples on bench top for at least 5 minutes at room temperature.
5m
Place the tube on an appropriate magnetic stand for 5 minutes to separate the beads from the supernatant.
Reminder: If necessary (i.e. not made earlier with other batch of 80% ethanol), prepare an additional ~360-400 µl of 80% freshly prepared ethanol per sample for the proceeding washes.
5m
After 5 minutes (or when the solution is clear), carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets (Caution: do not discard the beads).
1m
Add 200 μl of 80% freshly prepared ethanol to the tube while in the magnetic stand. Incubate at room temperature for 30 seconds, and then carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets.
1m
Repeat the 80% ethanol wash step once for a total of two washes. Be sure to remove all visible liquid after the second wash. If necessary, briefly spin the tube, place back on the magnet and remove traces of ethanol with a p10 pipette tip.
1m
Air dry the beads for up to 5 minutes while the tube/plate is on the magnetic stand with the lid open.
Caution: Do not over-dry the beads. This may result in lower recovery of DNA. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack, they are too dry.
5m
Remove the tube/plate from the magnetic stand. Elute the DNA target from the beads by adding 33 μl of 0.1X TE (dilute 1X TE Buffer 1:10 in water).
1m
Mix well on a vortex mixer. Incubate for at least 2 minutes at room temperature and ensure beads become resuspended in solution. Quickly spin the tube to collect the liquid from the sides before placing back on the magnetic stand.
2m
Place the tube on the magnetic stand. After 5 minutes (or when the solution is clear), transfer 30 μl of the final library to a new 0.2 ml tube and store at –20°C.
5m
Assess library quality and quantity
40m
Assess size distribution on an Agilent D1000 ScreenTape Assay for TapeStation Systems (4200) and determine library peak size. Ideal range between ~350-450 bp.
Requires 1 µl of library.
20m
Assess library concentration using the QuBit DNA BR Assay kit (or alternative qPCR-based quantification such as the KAPA Library Quantification Kits).
Requires 1-5 µl of library
20m
Final library pooling
Generate an appropriate pooling strategy for your libraries to be run across Illumina flowcell(s). Generally, pooling in equal molar concentrations is performed. To obtain the molar concentration (nM) for each library, perform the following calculation for each sample:

nM library =
QuBit DNA concentration ng/µL * 106
/
660 *​ TapeStation fragment size

(note 660 g/mol is the approximate molecular weight of 1 bp of dsDNA)

Then, calculate the volume of each sample library to add to achieve the desired per sample nM:

Pooling volume =
Desired nM concentration
/
nM

One technique is to adjust the Desired nM concentration up and down until the overall volume of your pool (including all libraries) is sufficient to be loaded on/across the desired flow cells and that the pooling volume per sample is at a feasible volume (i.e. between 2-10 µl).
Store final pooled libraries (of up to 480 individual libraries if using all 5 plates of a NEBNext Multiplex Oligos for Illumina 96 Unique Dual Index Primer Pairs Sets 1-5) at -20ºC until ready for sending to sequencing facility, or loading on the Illumina flowcell.
Protocol references
Urine filtration microscopy for S. haematobium egg detection performed as described in Knopp et al. BMC Infectious Diseases (2026)26:263.

Whole genome amplification adapted from illustra Ready-To-Go GenomiPhi V3 DNA Amplification Kit Product booklet (29018371 AC accessed 07.05.2026), and methods described in Shortt et al 2017, Le Clec’h et al 2018, and Nikolakis et al 2021.
Acknowledgements
The Whole Genome Amplification protocol was adapted from GenomiPhi V3 ‘DNA Amplification from Blood-blotted Paper’ protocol from the illustra Ready-To-Go GenomiPhi V3 DNA Amplification Kit Product booklet (29018371 AC accessed 07.05.2026), and methods described in Shortt et al 2017, Le Clec’h et al 2018, and Nikolakis et al 2021.