Schistosoma Species and Hybrid Genotyping With a Field Deployable Multi‐Marker Amplicon Panel

Abstract

Schistosomes are parasitic trematode worms of the genus Schistosoma, responsible for causing urogenital and intestinal schistosomiasis. Six primary species infect humans: S. mansoni, S. japonicum, S. mekongi, S. intercalatum, S. guineensis, and S. haematobium. In addition, several species including S. bovis, S. curassoni, and S. mattheei primarily infect animals, particularly cattle. Hybridization events have been documented both between human‐infecting species and between human‐ and animal‐infecting species of Schistosoma. Current methods for detecting hybrids rely on genotyping mitochondrial cytochrome oxidase I (cox1) and the ribosomal internal transcribed spacer (ITS), or whole‐genome sequencing. This protocol describes a multi‐locus, multiplex amplicon sequencing approach using Nanopore technology, referred to as NMAS‐Seq, for species‐level genotyping and hybrid detection. It includes comprehensive steps for multiplex nested PCR, amplicon quantification and pooling, library preparation, and sequencing. The workflow is adaptable for molecular typing of other parasitic species. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1: DNA extraction and multiplex nested PCR

Basic Protocol 2: Amplicon quantification and pooling

Basic Protocol 3: Amplicon library preparation and Nanopore sequencing

INTRODUCTION

According to the World Health Organization (WHO), schistosomiasis affects an estimated 240 million people worldwide, with an additional 700 million at risk in endemic areas (WHO, ^2025a). The disease primarily impacts school‐aged children in low‐resource communities lacking access to clean water and adequate sanitation, particularly in tropical and subtropical regions (WHO, ^2025b). In line with its global health agenda, the WHO has set a target to eliminate schistosomiasis as a public health problem by 2030 (WHO, ²⁰²⁰). However, significant gaps in epidemiological data present challenges in accurately mapping disease distribution and implementing effective intervention strategies.

Historically, Schistosoma species were considered to be host specific. Increasing evidence now shows that human‐infecting schistosomes can hybridize with other human‐ or animal‐infecting species, resulting in viable hybrids capable of infecting humans, animals, or both (Leger & Webster, 2017; Rey et al., ²⁰²¹). In Africa, frequent natural intra‐ and inter‐clade hybridization among Schistosoma species has reshaped the epidemiological landscape, bringing new attention to the zoonotic potential of these parasites (Leger & Webster, 2017; Panzner, ²⁰²¹). Hybridization can give rise to novel genotypes, potentially altering parasite virulence, host specificity, transmission dynamics, and drug susceptibility (King et al., 2015; Panzner, ²⁰²¹, Zacharia et al., 2022). Currently, hybrid detection relies mainly on mitochondrial cytochrome c oxidase subunit 1 (CO1) and the nuclear ribosomal internal transcribed spacer (ITS) markers. While useful, these markers have inherent limitations (King et al., 2015; Leger & Webster, ²⁰¹⁷; Panzner, ²⁰²¹; Rey et al., ²⁰²¹). CO1 is maternally inherited, it is only useful in the detection of hybrids when the mitochondrial haplotype differs from at least one nuclear marker. Likewise, the ITS marker is relatively conserved across Schistosoma species within the same clade, reducing its resolution for differentiating between closely related species. Similarly, existing microsatellite panels often depend on species‐specific primers, which fail in hybrids due to primer dropout (Lund et al., 2022). Although whole genome sequencing (WGS) offers the highest resolution for identifying hybrids, it remains expensive, technically demanding, and largely inaccessible in endemic settings.

Multi‐locus sequence typing (MLST), originally developed for bacterial genotyping, has successfully been applied to explore the population genetic structure and evolutionary history of various parasites (Cacciò et al., 2016; Costache et al., ²⁰²⁰; Gilchrist, ²⁰¹⁴; Lauthier et al., ²⁰¹²; Wang et al., ²⁰¹²; Yeo et al., ²⁰¹¹). The method generates genetic polymorphism data from multiple chromosomal regions, which are unlikely to be inherited together through a single genetic event. The increasing availability of reference genomes for parasitic species has made it easier to identify informative loci for MLST studies (Urwin & Maiden, 2003). When coupled with next‐generation sequencing technologies, MLST becomes a scalable and cost‐effective tool for genotyping and the detection of hybrids because it is capable of analyzing multiple loci and samples in parallel (Levitt et al., 2017).

This article introduces an alternative genotyping strategy called NMAS‐Seq, a multi‐locus, multiplex amplicon sequencing approach using Oxford Nanopore Technologies (ONT). NMAS‐Seq provides a low‐cost, high‐resolution alternative to conventional mitochondrial and nuclear marker genotyping strategies, enabling the ability to screen large numbers of schistosome miracidia with enhanced sensitivity for detecting species diversity and hybridization. The method utilizes 10 newly developed and validated pan‐genus markers, each containing single nucleotide polymorphisms (SNPs) that distinguish among human as well as animal Schistosoma species. The marker panel is particularly suited for resolving complex hybrid profiles, including distinguishing first‐generation hybrids from later‐generation recombinants where species contributions are no longer evenly split.

Basic Protocol 1 explains the process of DNA extraction and the preparation of primer mixes for multiplex‐nested PCR. Basic Protocol 2 describes the quantification of multiple amplicons, the calculation needed to pool concentrations of amplicons for multi‐locus sequencing success. Basic Protocol 3 describes the library preparation procedure utilized for Nanopore MinION sequencing.

CAUTION: Schistosoma spp. are human and animal pathogens. Live cercariae can penetrate skin of humans and animals causing infection. The organism is categorized as a Biosafety Level 2 (BSL‐2) pathogen. All handling of pathogenic organisms should follow appropriate guidelines and regulations.

Basic Protocol 1. DNA EXTRACTION AND MULTIPLEX NESTED PCR

This protocol outlines the procedures for DNA extraction and two rounds of PCR amplification for multilocus genotyping. The first round consists of a multiplex PCR using two external primer mixes (PMIX 1 and PMIX 2). The second round involves nested PCR, where individual gene markers are amplified using their respective internal primers.

Strategic planning

This protocol outlines the procedures for parasite storage, selection of gene markers for multilocus sequence typing (MLST), and the grouping of these markers for multiplex PCR applications.

Storage of parasite

This method enables multilocus genotyping of Schistosoma larval stages directly from field‐collected samples, such as individual Schistosoma eggs hatched into miracidia or cercariae stored on Whatman FTA cards (Gower et al., 2013). The same approach is also compatible with parasite stages preserved in RNA Later or 70% ethanol.

Gene marker selection

This technique involves the amplification and sequencing of multiple enzyme‐coding, housekeeping, and protein‐coding genes. In total, nine nuclear genes and one mitochondrial gene were selected, in addition to the routinely used mitochondrial cytochrome oxidase subunit 1 (CO1) and the nuclear internal transcribed spacer (ITS) region. The selection criteria included: (a) single‐copy gene status, (b) easy sequence alignment across multiple species, (c) feasibility for pan‐genus primer design, (d) amplification efficiency, (e) informative sequence variation, and (f) orthology (Lauthier et al., 2012; Wang et al., ²⁰¹²). The ten selected gene targets span six chromosomes in the S. haematobium reference genome. The nonsynonymous to synonymous substitution ratio (dN/dS) was calculated using the Nei‐Gojobori method via SNAP software (http://www.hiv.lanl.gov) (Nei & Gojobori, 1986). Primers were designed to amplify fragments ranging from 279 to 600 bp. Detailed information on marker genes, chromosomal locations, primer sequences, and fragment lengths is provided in Table 1.

Table 1.

Selected Gene Markers

Chr a	Gene	Gene ID	External primers	Sequence	Length (Ext)	Internal primers	Sequence	Length (Int)
1	Sel‐1‐like protein	SP	SPEXF	YGATGATCCGGAAACTGTCG	753	SPINF	AACTGCGAACGATTTTCCGG	453
			SPEXR	TAGACAGCTCTTCGAACGCA		SPINR	AACCTCCAAGCAATGTCAGC
1	ATP‐binding cassette	AT	ATEXF	CCCTTCTGGCGTGGAAATTT	456	ATINF	AAAACGTGGTCAGTGGTGGT	279
			ATEXR	GAATCCAGCCAACACACCAG		ATINR	CCTCGGAGTATGTTTGCACC
3	DUF2040 domain‐containing protein	DUF	DUFEXF	GCGGTAAGGTGTACGGYCTT	596	DUFINF	GAACTTGTAGGGGTTTCATCGT	371
			DUFEXR	TCTCCTCTTCTTCCTTGCGT		DUFINR	TTCCTTCTGAGCCTTCCGTT
4	T‐complex protein 1 subunit gamma	TSG	TSGEXF	GYTTAGGWCCWAGGGCAATG	700	TSGINF	TGCTCATGGATCCTATGGGT	554
			TSGEXR	AACGCCATCAAGAACAAYYG		TSGINR	CCGTCGGTTTCCACTGTTAC
5	Transaldolase	T	TEXF	TYYCAAATCTTTCAGTYMCTTC	633	TINF	GTAGCGGACACTGGAGATTT	363
			TEXR	GCRGCTTTTATACCTTCCCATGT		TINR	ACCGAGCATCCACTTCAGTA
7	Bravo_FIGEY domain‐containing protein	BF	BFEXF	TGTATCACGCTGGCCATACT	712	BFINF	ACTAGATGGCAGATACGGACC	408
			BFEXR	CCACCTGCCATCAAACTCAC		BFINR	TAGTCCCCTTGAGGTTGTCG
7	Putative tropomyosin	TY	TYEXF	TCCAGTGGAATTACTCGGACC	730	TYINF	TCGATGACAACTCTGCTATGGA	456
			TYEXR	TCATCAGCAGCTTTACTCGC		TYINR	TCTCTTCTGTAGTGAGGCGAC
ZW	ADP/ATP translocase	ATP	ATPEXF	ACACATGAACGTTTACTAGAGGT	729	ATPINF	GCAAAAGCAGCCGAACTTCT	565
			ATPEXR	GGTCAATTTCTGCTTGTTCATCA		ATPINR	CCAAACGCATGCATTCAAATCC
ZW	60S ribosomal protein L8	RPL	RPLEXF	GGTTATTCGWAGYCARCGTA	550	RPLINF	GGAGGTGTTTTCAAAGCGCA	502
			RPLEXR	CTGACTGTGGGCCAACAATG		RPLINR	ACAAGACCTACCATGGCACG
ML	Internal transcribed spacer 2	ITS	ITSEXF	GTGCAGCCAACTGTGTGAAT	600	ITSINF	CATATTGCGGCTACGGGATA	398
			ITSEXR	TTGGGCTAATCCCTGTTCAC		ITSINR	AAGTTCAGCGGGTAATCACG
MG	Cytochrome oxidase subunit 1	CO	COEXF	TGGTTGTGGTGTAGGATGAACA	598	COINF	GCGGGTGTATCTAGACTAGTTGG	530
			COEXR	ATACCAGTAACACCACCTATCGT		COINR	ACACGAGACCCACAGCTTTT
MG	NADH dehydrogenase subunit 5	ND	NDEXF	GGGTAAAAGTTGGAATTTGAGGG	625	NDINF	GTCTTATCGTGGTGGAGTGGT	459
			NDEXR	CGCTTTAACCATCTGACCACC		NDINR	TCCACTCTCCCCATAACCATCA

^a ML = multiple loci, MG = mitochondrial genome.

Grouping of gene markers

To facilitate multiplex PCR, gene markers were divided into two groups based on fragment size and external primer sequence dissimilarity.

Group 1 includes primers (purchased from Integrated DNA Technologies) targeting the following genes:

• Sel‐1‐like protein

• ATP‐binding cassette

• T‐complex protein 1 subunit gamma

• NADH dehydrogenase subunit 5

• DUF2040 domain‐containing protein

• ADP/ATP translocase.

Group 2 includes primers for:

• Putative tropomyosin

• Bravo_FIGEY domain‐containing protein

• Transaldolase

• Cytochrome oxidase subunit 1 (CO1)

• 60S ribosomal protein L8

• Internal transcribed spacer 2 (ITS2).

Preparation of external primer stocks for multiplex PCR

• Centrifuge all external primers briefly.

• Label two microcentrifuge tubes as PMIX 1 (Group 1) and PMIX 2 (Group 2).

• For PMIX 1: From 100 µM stock solutions, prepare a 2 µM multiplex mix by combining 10 µl each of the forward and reverse primers for all six Group 1 genes (12 primers total), and add 380 µl of nuclease‐free water.

• For PMIX 2, repeat the same procedure using the Group 2 primers.

• Mix thoroughly by pipetting and briefly centrifuge to collect contents.

• Store both primer mixes at –20°C.

Additional planning

• Label PCR tubes with both sample ID and primer name to ensure traceability.

• Limit processing to no more than 20 samples at a time to reduce the risk of pipetting or labeling errors.

• Use multichannel pipettes where possible to streamline workflows and reduce pipetting time.

• Include a negative control (no template DNA) in both PCR rounds to monitor for contamination.

Materials

• Whatman FTA cards (Sigma, WHAWB120205 or WHA10534612) or worms/eggs stored in RNAlater stabilization solution (Thermo Fisher, AM7020) or 70% ethanol

• DNeasy Blood and Tissue Kit (Qiagen, 69506) containing:

  • Buffer ATL

  • Proteinase K

  • Buffer AL

  • Buffer AW1 concentrate

  • Buffer AW2 concentrate

  • Mini Spin columns

  • 2‐ml collection tubes

• 100% ethanol

• H₂O, Ambion nuclease‐free (Thermo Fisher, AM9937)

• PMIX 1 (see Strategic planning above)

• PMIX 2 (see Strategic planning above)

• 5× KAPA HiFi buffer (Roche, 07958846001):

  • 10 mM KAPA dNTP mix

  • 1 U/µl KAPA HiFi DNA ploymerase

• Gel electrophoresis reagents:

  • UltraPure agarose powder (Thermo Fisher, 16500‐900)

  • TAE/TBE (Apex Bioresearch, 20‐194)

  • GelRed stain (VWR, 89139‐142)

  • Bromophenol blue tracking dye (Quality Biological, 351‐028‐661)

• Unicore punch kit, 2.0‐mm (Qiagen, WB100029)

• 1.5‐ml Maxymum Recovery Snaplock microcentrifuge tubes (Axygen, MCT‐150‐L‐C)

• Pipettes (Gilson and Rainin brands)

• 10‐, 200‐, and 1000‐µl filtered pipette tips (PurePoint extra‐long, FT1010; PurePoint, FT1200; and CellTreat low retention extended length, 229022)

• Vortex mixer

• Incubator

• Eppendorf centrifuges 5810R and 5430R

• TempAssure 0.2‐ml PCR 8‐tube strips with individual caps attached (USA Scientific, 1402‐4700)

• Eppendorf Mastercycler nexus thermal cycler

• Gel electrophoresis equipment

• UV or blue‐light transilluminator

• Multichannel pipettes (Thermo Fisher, Finnpipette)

DNA extraction

Genomic DNA is extracted directly from Whatman FTA cards using the Qiagen DNeasy Blood and Tissue Kit following the manufacturer's instructions. Extracted DNA can be used immediately or stored at –20°C for later use.

• 1Prepare samples. Using a 2‐mm punch, cut out individual samples from Whatman FTA cards and place each into a 1.5‐ml microcentrifuge tube.
• 2Lysis step. Add 180 µl Buffer ATL and 20 µl proteinase K to the tube. Mix thoroughly using a vortex mixer, then incubate at 56°C for 2 hr.
• 3Cell disruption and preparation for binding. Vortex the tube for 15 s and add 200 µl Buffer AL and mix thoroughly by vortexing. Add 200 µl of 100% ethanol and mix again by vortexing.
• 4DNA binding. Transfer the entire mixture (including any precipitate) to a DNeasy Mini spin column placed in a 2‐ml collection tube. Centrifuge 1 min at ≥6000 × g, room temperature. Discard the flowthrough along with the collection tube.
• 5Wash Step 1. Place the spin column into a new 2‐ml collection tube. Add 500 µl Buffer AW1, centrifuge 1 min at ≥6000 × g, room temperature, and discard the flowthrough and tube.
• 6Wash Step 2. Place the Mini spin column into a fresh 2‐ml collection tube. Add 500 µl Buffer AW2, centrifuge 3 min at ≥20000 × g, room temperature, and discard the flowthrough.
• 7Dry spin. Return the spin column to the same collection tube and centrifuge 1 min at ≥20000 × g, room temperature, to remove residual wash buffer.
• 8Elution. Transfer the Mini spin column to a clean 1.5‐ml microcentrifuge tube. Add 10 µl of nuclease‐free water directly to the membrane, incubate at room temperature for 1 min, and centrifuge 1 min at ≥6000 × g, room temperature. Repeat the elution step with an additional 5 µl of nuclease‐free water to obtain a total volume of 15 µl of eluted DNA.
• 9Store DNA samples at 4°C for short‐term use or at –20°C for long‐term storage.

PCR 1: Multiplex PCR using PMIX1 and PMIX2 primers

For each DNA sample, perform two separate multiplex PCR reactions: one with PMIX1 and one with PMIX2 primer mixes. Follow the steps below to prepare reactions and run the thermal cycling protocol.

• 10Prepare master mixes. Prepare separate master mixes for PMIX1 and PMIX2 reactions according to the reagent volumes listed in Table 2.

  Multiply the volumes by the number of reactions and include at least 10% extra volume to compensate for pipetting loss.

Table 2.

PCR Reagents for 25 µl Single Reaction

Reagent	Volume (µl)	Final concentration
Nuclease‐free water	13
KAPA HiFi Buffer	5	1×
1 U/µl KAPA HiFi DNA polymerase	0.5 µl	0.5 U
10 mM KAPA dNTP Mix	0.75	3 mM
Primer	3.75	0.5 µM
Template DNA	2ul

• 11Aliquot master mix. Dispense 23 µl of the appropriate master mix into each labeled PCR tube (label with sample ID and primer group: PMIX1 or PMIX2).
• 12Add template DNA. Add 2 µl of extracted genomic DNA to each tube to bring the total reaction volume to 25 µl.
• 13Thermal cycling. Load the PCR tubes into a thermal cycler and run the PCR using the cycling conditions specified in Table 3.

  PCR‐strip tubes or 96‐well plates are recommended for the PCR reactions.

Table 3.

PCR Cycling Conditions

No. of cycles	Time	Temperature	Purpose
1 cycle	3 min	95°C	Initial denaturation
35 cycles	20 s	98°C	Denaturation
	15 s	58°C	Annealing
	30 s	72°C	Extension
1 cycle	10 min	72°C	Final extension

Gel electrophoresis (optional)

This optional step provides a brief overview of gel electrophoresis to check PCR amplification (Fig. 1). Note that bands may not be visible for all loci.

Figure 1.

Gel Image for multiplex PCR performed on DNA extracted from schistosome ova‐positive animal and patient samples. SH = S. haematobium, SM = S. mansoni, SB = S. bovis, NC = negative control.

• 14Prepare agarose gel. Dissolve 3 g agarose in 100 ml of 1× TBE or TAE buffer to make a 0.3% agarose gel. Heat until fully dissolved.
• 15Add DNA stain. Once the agarose has cooled to ∼60°C, add 4 µl of GelRed (or an equivalent nucleic acid stain), and mix thoroughly before pouring into the gel tray with combs in place.
• 16Prepare samples for loading. Mix 4 µl of PCR product with 1 µl loading dye. Load into wells of the solidified gel.
• 17Run gel. Run the gel at 110 V for 30 to 35 min in 1× TBE or TAE running buffer.
• 18Visualize DNA bands using a UV or blue‐light transilluminator.

PCR II: Nested PCR using individual internal primers

For each DNA sample, perform independent PCR reactions with each of the 12 internal primer pairs to amplify specific gene targets. This step follows the initial multiplex PCR (PCR I) and uses the product from that reaction as template.

• 19Prepare master mixes. For each internal primer pair, prepare a separate master mix based on the reagent volumes listed in Table 4.

  Multiply the volumes by the number of reactions and include at least 10% extra volume to compensate for pipetting loss.

Table 4.

PCR Reagents for 25 µl Single Reaction

Reagent	Volume (µl)	Final concentration
Nuclease‐free water	15.25
KAPA HiFi Buffer	5	1×
1 U/µl KAPA HiFi DNA polymerase	0.5 µl	0.5 U
10 mM KAPA dNTP Mix	0.75	3 mM
Forward Primer	1.25
Reverse Primer	1.25	0.5 µM
Amplified DNA (from PCR 1)	1

• 20Aliquot master mix. Dispense 24 µl of the appropriate master mix into labeled PCR tubes (include sample ID and primer name).

  Use a multichannel pipette, if possible, to speed up processing.

• 21Add PCR template. Add 1 µl of the corresponding PCR I product (from PMIX1 or PMIX2) to each tube, bringing the final volume to 25 µl.
• 22Thermal cycling. Load tubes into the thermal cycler and run the PCR using the cycling conditions specified in Table 3.

Gel electrophoresis

This step provides a brief overview of gel electrophoresis to check PCR amplification. Conduct workflow as described above after PCR 1. Note that bands should be visible for all loci (Fig. 2).

Figure 2.

Gel Image for singleplex PCR for all markers using genomic DNA. Although the bands for CO1 and ND were less prominent for S. mansoni, the PCR worked well even in field samples. SH = S. haematobium, SM = S. mansoni, SB = S. bovis, NC = negative control.

Basic Protocol 2. AMPLICON QUANTIFICATION AND POOLING

This protocol outlines the steps for quantifying PCR amplicons and pooling gene loci for each sample in preparation for sequencing.

Strategic planning

Organize PCR products according to their respective gene loci in preparation for barcoding and sequencing. Refer to the Oxford Nanopore Technologies Native Barcoding Kit 24 V14 (SQK‐NBD114.24) or Native Barcoding Kit 96 V14 (SQK‐NBD114.96) protocol for detailed instructions on barcoding and library preparation. ONT recommends using 200 fmol of DNA per sample for barcoding. For 1 kb amplicons, this corresponds to ∼130 ng of DNA. To determine the DNA mass equivalent to 200 fmol for each amplicon based on fragment length, calculations were performed using the Promega Biomath Calculator (see Internet Resources). The required DNA quantities for each gene locus are summarized in Table 5.

Table 5.

DNA Concentration (ng) Equivalent to 200 fmol for Each Gene

Chr a	Gene ID	Length (Int)	Concentration (ng)
1	SP	453	60
	ATP	279	37
3	DUFF	371	49
4	TSG	554	73
5	T	363	48
7	BF	408	54
	TY	456	60
ZW	AT	565	75
	RPL	502	66
ML	ITS	398	53
MG MG	ND5 COX	459 530	61 70
	Total Size	5338 bp

^a ML = multiple loci, MG = mitochondrial genome.

CAUTION: Do not use a NanoDrop spectrophotometer for DNA quantification, as it lacks the sensitivity and accuracy required for low‐concentration amplicons. Use a fluorometric method (e.g., Qubit, or Promega QuantiFluor dsDNA System ) for reliable results.

Additional Materials (also see Basic Protocol 1)

• Qubit dsDNA BR assay kits (Thermo Fisher, Q32853) or QuantiFluor ONE dsDNA System (Promega, E4870)

• Fluorometer, e.g., Qubit 4 Fluorometer (Invitrogen, Q33226) or Quantus (Promega, E6150)

• Qubit 0.5‐ml assay tubes (Thermo Fisher, Q32856)

• Mini centrifuge

• 1Using a fluorometer, assay tubes, and the Qubit dsDNA BR assay kits or QuantiFluor ONE dsDNA System assay kits, quantify five randomly selected samples for each gene locus.

  Selective random quantification is recommended to manage workload when processing a large number of samples.

• 2Calculate the average DNA concentration for each gene locus and determine the volume of amplified DNA needed for pooling.

  For example, for the gene marker SP amplified with primers SPINF and SPINR, 60 ng of DNA corresponds to 200 fmol. If the average DNA concentration from step 1 is 90 ng/µl, the volume needed for pooling is ∼0.7 µl (calculated as 60 ng ÷ 90 ng/µl).

• 3Repeat steps 1 and 2 for all gene loci to determine the required pooling volume for each locus.
• 4Sum the individual volumes to calculate the total expected volume for each sample pool.
• 5Label new PCR tubes clearly with the corresponding sample IDs.
• 6Pool the calculated volumes of all gene loci for each individual sample into the corresponding labeled tube.
• 7Mix gently by pipetting up and down, then briefly spin down using a mini centrifuge at maximum speed, to collect the liquid at the bottom of the tube.
• 8The total number of pools depends on the Oxford Nanopore Technologies Native Barcoding Kit used.

  For example, with a 24‐barcode kit, pool 22 samples along with two negative controls.

Basic Protocol 3. AMPLICON LIBRARY PREPARATION AND SEQUENCING

This protocol outlines the workflow for library preparation and multiplexed amplicon sequencing using Oxford Nanopore Technologies. Sequencing will be performed on the MinION platform following ONT's standard procedures.

Strategic planning

Determine read number for sequencing coverage

Calculate the minimum number of reads per barcode required to achieve the desired sequencing coverage. Consider important factors, such as the total size of the pooled loci (in base pairs), read filtering efficiency, and mapping efficiency. This calculation is essential for estimating the duration of the sequencing run. The number of reads needed can be estimated using the formula:

$$\mathrm{Number}\mathrm{of}\mathrm{reads}=\frac{\mathrm{Average}\mathrm{read}\mathrm{length}}{\mathrm{Target}\mathrm{coverage}\times \mathrm{Amplicon}\mathrm{length}}$$

For example, Average read length = 5338 bp, Target coverage = 100×, and Amplicon length = 5338 bp.

$$\mathrm{Number}\mathrm{of}\mathrm{reads}=\frac{5338}{100\times 5338}=100\mathrm{reads}\mathrm{per}\mathrm{barcode}$$

Assume 40% uneven barcode distribution (i.e., effective reads = 60%), 50% of reads filtered out during quality control, and 50% mapping efficiency.

$$\mathrm{Number}\mathrm{of}\mathrm{reads}=\frac{100}{0.6\times 0.5\times 0.5}=667\mathrm{reads}\mathrm{per}\mathrm{barcode}$$

Thus, a minimum of 667 reads per barcode is required to achieve ∼100× coverage.

Determine pool concentration

Calculate the concentration of your pooled amplicon sample using:

$$\mathrm{Pool}\mathrm{concentration}(\mathrm{y})=\frac{\mathrm{Total}\mathrm{concentration}}{\mathrm{Total}\mathrm{volume}(\mathrm{x})}=\frac{706\mathrm{ng}}{\mathrm{x}\mu \mathrm{l}}$$

The total volume × is obtained from Basic Protocol 2, step 4.

Determine volume for sequencing

Average gene length ∼445 bp. Using the Promega Biomath calculator (see Internet Resources), the DNA mass corresponding to 200 fmol for 445 bp is ∼58 ng. Calculate the volume v of pooled DNA to use for sequencing based on pool concentration y.

$$\mathrm{Volume}\mathrm{for}\mathrm{sequencing}(\mathrm{v})=\frac{58\mathrm{ng}}{\mathrm{y}}$$

Additional Materials (also see Basic Protocol 1)

• Oxford Nanopore Technologies Native Barcoding Kit 24 V14 (SQK‐NBD114.24) or Native Barcoding Kit 96 V14 (SQK‐NBD114.96):

  • Native Barcode plate (NB01‐24 or NB01‐96)

  • DNA Control Sample (DCS)

  • Native Adapter (NA)

  • Sequencing Buffer (SB)

  • Library Beads (LIB)

  • Library Solution (LIS)

  • Elution Buffer (EB)

  • AMPure XP Beads (AXP)

  • Long Fragment Buffer (LFB)

  • Short Fragment Buffer (SFB)

  • EDTA

  • Flow Cell Flush (FCF)

  • Flow Cell Tether (FCT)

• New England Biolabs (NEB) specific reagents for Oxford Nanopore Technologies:

  • NEBNext Ultra II End Repair/dA‐Tailing module (NEB, E7546)

  • NEB Blunt/TA Ligase master mix (NEB, M0367)

  • NEBNext Quick Ligation module (NEB, E6056)

• 200 fmol (130 ng for 1 kb amplicons) DNA per sample to be barcoded

• Qubit dsDNA HS assay kit (Thermo Fisher, Q32851)

• Ice bucket with ice

• Mini centrifuge

• 0.2‐ml non‐skirted low profile 96‐well PCR plate (Thermo Scientific, AB‐700)

• Microseal “B” seals (BioRad, MSB1001)

• Microplate centrifuge

• HulaMixer, i.e., gentle rotator mixer

• Invitrogen DynaMag‐96 Side (Thermo Fisher, 12331D)

• Qubit assay tubes (Invitrogen, Q32856)

• MinION Mk1B (Oxford Nanopore Technologies)

• R10.4.1 flow cells (Oxford Nanopore Technologies, FLO‐MIN114)

End‐prep

This step will prepare the DNA ends for attachment of sequencing adapter.

• 1Thaw reagents. Thaw the DNA Control Sample (DCS) at room temperature. Mix thoroughly by vortexing, then place on ice. Thaw the NEBNext Ultra II End Repair reagents on ice. Mix gently by pipetting and briefly spin down using a mini centrifuge at maximum speed at room temperature to collect contents.
• 2Dilute the Control DNA. Add 105 µl Elution Buffer (EB) directly to one tube of the DNA Control Sample (DCS). Mix gently by pipetting and spin down at room temperature. Keep on ice until use.
• 3Aliquot sample DNA. In a clean 96‐well plate, aliquot the calculated volume of DNA for each sample (refer to Strategic planning above for guidance). Adjust each sample to a final volume of 11.5 µl using nuclease‐free water. Mix gently by pipetting and spin down at room temperature.
• 4Prepare End‐Prep master mix. Prepare a master mix based on Table 6, scaling to the number of samples being processed. Include extra volume to account for pipetting loss.

Table 6.

Preparation of End‐Prep Master Mix

Reagent	Volume (µl)
Ultra II End‐prep reaction buffer	1.75
Ultra II End‐prep enzyme mix	0.75
Total	2.5

• 5Add reagents to samples. Add 2.5 µl of End‐Prep master mix to each well. Add 1 µl of the diluted DNA Control Sample (DCS) to each sample well. Mix thoroughly by pipetting, seal the plate and briefly spin down in a microplate centrifuge at room temperature.
• 6Incubation. Place the plate or tubes in a thermal cycler and incubate at 20°C for 5 min and 65°C for 5 min.
• 7Hold at 4°C or place on ice until proceeding to adapter ligation.

Native barcode ligation

This step describes the ligation of native barcodes to end‐prepped DNA using the Oxford Nanopore Technologies barcoding workflow. Each sample receives a unique barcode to allow for multiplexed sequencing.

• 8
  Thaw the following reagents at room temperature:

  • NEB Blunt/TA Ligase Master Mix
  • AMPure XP Beads
  • EDTA
  • Native Barcodes (thaw only the number needed, one per sample).
    Up to 96 samples can be barcoded and pooled in a single run.
• 9Mix the NEB Blunt/TA Ligase master mix. Spin down 5 s at room temperature. Mix thoroughly by pipetting up and down 10 times using the full volume.
• 10Vortex the AMPure XP Beads and keep at room temperature.
• 11Vortex the EDTA, spin down, and place on ice.
• 12Mix each barcode (selected for the samples) by pipetting, spin down briefly, and place on ice.
• 13In a clean 96‐well plate, add the reagents in the order listed to each well, mixing gently by pipetting after each addition (Table 7)

Table 7.

Preparation of Adapter Ligation Reaction

Reagent	Volume (µl)
Nuclease‐free water	3
End‐prepped DNA	0.75
Native Barcode	1.25
Blunt/TA Ligase Master Mix	5
Total	10

• 14Mix thoroughly by pipetting and spin down briefly.
• 15Incubate at room temperature for 20 min.
• 16Add 1 to 2 µl EDTA to each well to stop the ligation reaction.

  Refer to the barcode kit manual to confirm the recommended EDTA volume.

• 17Mix thoroughly and spin down briefly.
• 18Pool all barcoded samples into a single 1.5‐ml microcentrifuge tube.

  Check the base of the plate to ensure complete transfer.

• 19Resuspend the AMPure XP Beads by vortexing.
• 20Add 0.4× AMPure XP Beads to the pooled sample and mix thoroughly by pipetting.
• 21Incubate on a HulaMixer at room temperature for 10 min.
• 22Spin down briefly, and place the tube on a magnetic rack for 5 min.
• 23Once the solution becomes clear and colorless, carefully remove and discard the supernatant.
• 24With the tube still on the magnetic rack, wash the beads with 700 µl of freshly prepared 80% ethanol without disturbing the pellet.
• 25Remove the ethanol carefully using a pipette.

  If the pellet is disturbed, wait for it to re‐pellet before proceeding.

• 26Repeat the ethanol wash (steps 23 to 24).
• 27Spin down briefly, and return the tube to the magnetic rack.
• 28Remove any residual ethanol with a fine pipette tip and let the pellet air‐dry for ∼30 s.

  Do not over‐dry the pellet to avoid cracking of the pellets.

• 29Remove the tube from the magnetic rack and resuspend the pellet in 35 µl of nuclease‐free water by gently flicking the tube.
• 30Incubate at 37°C for 10 min, gently flicking the tube every 2 min for 10 s to encourage elution.
• 31Return the tube to the magnetic rack and wait until the eluate is clear and colorless.
• 32Carefully transfer 35 µl of the eluate into a clean 1.5‐ml microcentrifuge tube.
• 33Quantify 1 µl of the eluted DNA using a fluorometer.

• 34Thaw the NEBNext Quick Ligation Reaction Buffer, Quick T4 DNA Ligase, and Native Adapter at room temperature.

  Do NOT vortex the Quick T4 DNA Ligase.

• 35Spin down briefly, then mix by pipetting and place the reagents on ice.

  If the Quick Ligation Reaction Buffer appears precipitated, pipette up and down several times to dissolve the precipitate, then vortex briefly to ensure thorough mixing.

• 36Thaw the Elution Buffer at room temperature. Vortex briefly, spin down, and place on ice.
• 37Thaw the Short Fragment Buffer at room temperature. Vortex briefly and spin down.

  The Short Fragment Buffer retains DNA fragments of all sizes.

• 38Prepare the adapter ligation reaction in a 1.5‐ml microcentrifuge tube (Table 8). Mix thoroughly by pipetting 10 to 20 times between each addition.

Table 8.

Preparation of Adapter Ligation Reaction

Reagents	Volume (µl)
Pooled barcoded sample	30
Native Adapter	5
NEBNext Quick Ligation reaction buffer	10
Quick T4 DNA Ligase	5
Total	50

• 39Mix the final reaction gently by pipetting and spin down briefly.
• 40Incubate the reaction for 20 min at room temperature.
• 41Resuspend the AMPure XP Beads by vortexing.
• 42Add 20 µl of resuspended AMPure XP Beads to the ligation reaction and mix by pipetting.
• 43Incubate on a HulaMixer for 10 min at room temperature.
• 44Spin down, and place the tube on a magnetic rack until the beads are fully pelleted.
• 45While the tube remains on the magnet, carefully remove and discard the supernatant.
• 46Wash the beads by adding 125 µl of Short Fragment Buffer.
• 47Flick the tube to resuspend the beads, spin down, and place it back on the magnetic rack to pellet the beads.
• 48Remove and discard the supernatant.
• 49Repeat the steps 46 to 48 for a second wash.
• 50Spin down, and return it to the magnetic rack.
• 51Remove any residual supernatant with a pipette.
• 52Remove the tube from the magnetic rack and resuspend the bead pellet in 15 µl of Elution Buffer.
• 53Spin down, and incubate for 10 min at 37°C, gently flicking or agitating every 2 min.
• 54Place the tube on the magnetic rack and allow the beads to pellet. Wait until the eluate is clear and colorless (at least 1 min).
• 55Carefully transfer 15 µl of the eluate (which contains the DNA library) to a clean 1.5‐ml microcentrifuge tube.
• 56Dispose of the pelleted beads.
• 57Quantify 1 µl of eluted sample using a Qubit fluorometer.
• 58Prepare the final DNA library, aiming for 35 to 50 fmol (120 to 180 ng) in 12 µl of Elution Buffer.
• 59Store the library on ice or at 4°C until ready to load.
• 60Follow the appropriate preparation for priming and loading the SpotON flow cell.

COMMENTARY

Background Information

Schistosomiasis is a neglected parasitic disease prevalent in Africa infecting both humans and animals. An estimated 700 million people, living in poor, rural communities without access to basic amenities, such as potable water and adequate sanitation, are at risk of various forms of schistosomiasis (Aula et al., 2021). The disease is transmitted by intermediate snail vector hosts that shed Schistosoma infective stages in freshwater bodies. Humans and animals contract the disease from water contact with infested water bodies where the infective stages of the parasite penetrate the human or animal hosts. Three Schistosoma species (S. mansoni, S. japonicum, and S. haematobium) are the widely recognized and studied causative agents of human schistosomiasis. S. japonicum and S. mansoni are responsible for intestinal schistosomiasis, while S. haematobium is responsible for urogenital schistosomiasis (WHO, ^2025b). In schistosomiasis infections, the adult stages of the parasites lodge in the host's blood vessels within the intestinal or urogenital systems, where the mature female produces thousands of eggs, which are deposited in various tissues and organs, most notably the liver, bladder, small and large intestine, cervix, and vagina. Most of the disease pathology arises from inflammatory reactions and immune responses due to trapped eggs in the various organs and can result in complications, such as hepatosplenomegaly, periportal fibrosis, bladder cancer, and female genital schistosomiasis (Colley et al., 2014; McManus et al., ²⁰¹⁸). Schistosomiasis infection is also linked to stunting and decreased cognitive performance in children, increased risk of HIV infection in women, ectopic pregnancies, and other long‐term irreversible consequences such as infertility (Aula et al., 2021).

Formerly, some schistosome species were thought to be host‐specific; however, advanced applications of molecular techniques have provided evidence that human and animal schistosomes can pair to produce zoonotic hybrids capable of infecting both humans and animals. In West Africa, hybrids of S. haematobium and livestock schistosomes, S. bovis and/or S. curassoni has been reported as evidence of possible zoonotic transmission of the parasite (Rey et al., 2021). Recent research on epidemiology of schistosomiasis is being focused on single egg genotyping and hybrid detection. The addition of new genetic markers and a field‐deployable sequencing platform should prove instrumental in clarifying the degree to which intra‐specific (within a species) and inter‐specific (between species) hybridization is occurring, which has important implications in host range expansion, disease, and transmission of this highly prevalent trematode.

Critical Parameters

The initial starting material contains ∼2 ng/µl of DNA; therefore, samples should be handled carefully to avoid DNA loss. All PCRs and library preparations should be performed in a biosafety cabinet to minimize environmental contamination. When conducting PCR, negative controls must be included. For library preparation, both negative and positive controls may be included. The positive control can be sourced as genomic DNA from BEI Resources or as adult worms from the Schistosomiasis Collection at the Natural History Museum (SCAN), UK.

The DNA extraction kits and PCR amplification reagents described above successfully isolated Schistosoma DNA and produced robust PCR amplifications. Other kits should not be used without proper evaluation. Other high‐fidelity Taq reagents may be assessed for performance. If inhibitors are suspected in the PCR reaction, bovine serum albumin (BSA) can be added.

During gel electrophoresis, double bands were observed at the recommended annealing temperature of 58°C. However, the presence of double bands does not affect the sequencing reaction, if Schistosoma DNA is confirmed at the correct band size for the targeted markers.

The sequencing should be run depending on the desired read depth. It takes ∼6 hr to obtain a read depth of 100× for at least 90% of a 96‐barcodes run.

Troubleshooting

These are straightforward protocols with minimal issues. The commonly encountered problems and possible solutions are shown in Table 9.

Table 9.

Troubleshooting Guide for PCR, Library Prep, and Sequencing

Problem	Possible cause	Solution
No amplification	No or very low concentration of DNA	Use Qubit fluorometer to detect DNA concentration
	No parasite sample loaded in FTA card	Review extraction kit and protocol
	Primers degraded	Prepare fresh dilutions of primers from stock solutions
Faint gel bands	Very low concentration of DNA	Determine concentration using Qubit fluorometer and calculate pooling volume as recommended
Low number of reads	Poor starting material	Quantify samples pre‐ and post‐pooling
	Short sequencing run	Sequence for longer period

Understanding Results

Gel electrophoresis images are useful for confirming amplification of target loci and pooling prior to sequencing (Fig. 2). The ONT sequencing platform outputs demultiplexed FASTQ files for each sample based on the number of barcodes. The data analysis follows the recommended NGS data pipeline for variant calling analysis. A curated reference genome for S. haematobium can be found in the supplementary file (see Supporting Information). Depending on the analysis, reference genome can be curated for other Schistosoma species. Software, e.g., NanoFilt, Minimap2, Samtools, and BCFtools, recommended for analysing ONT data, should be used for read filtering, mapping, and generating SNP data. BAM files can be visualized in IGV. SNP data can be further analysed and visualized by generating consensus sequence files in FASTA format. These can be imported into Geneious for multiple sequence alignments. Phylogenetic data analysis can be conducted in MEGA X using the maximum likelihood method with 1000 bootstraps. Potential outputs include allelic plot, phylogenetic trees, and haplotype networks.

Time Considerations

The PCR depends on the number of samples being processed at once. Each round of PCR takes 1.5 hr. Pooling and library preparation usually takes ∼6 hr.

Author Contributions

Oluwaremilekun G. Ajakaye: Conceptualization; investigation; writing—original draft; writing—review and editing. Michael E. Grigg: Funding acquisition; project administration; supervision; writing—review and editing.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Excel file containing curated reference genome for S. haematobium.

Ajakaye, O. G. , & Grigg, M. E. (2026). Schistosoma species and hybrid genotyping with a field deployable multi‐marker amplicon panel. Current Protocols, 6, e70333. doi: 10.1002/cpz1.70333

Data Availability Statement

All information required to run the protocol is included in the manuscript.