A Recombinase Polymerase Amplification Lateral Flow Assay (RPA-LF) for detection of Babesia divergens

A Recombinase Polymerase Amplification Lateral Flow Assay (RPA-LF) for detection of Babesia divergens | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article A Recombinase Polymerase Amplification Lateral Flow Assay (RPA-LF) for detection of Babesia divergens Ernest Chin Jia Bin, Keir Hughes, Cheryl Whitehorn, Ellen knuepfer, and 6 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8999595/v1 This work is licensed under a CC BY 4.0 License Status: Under Revision Version 1 posted 9 You are reading this latest preprint version Abstract Background: Babesia divergens is a tick-borne apicomplexan parasite found predominantly in Europe, causing babesiosis in livestock and occasionally in humans, with severe disease reported in immunosuppressed or asplenic individuals. As populations of Ixodes ricinus , the principal vector, expand geographically including into urban environments, there is a growing need for rapid, sensitive, and field-deployable surveillance tools. In this study, we developed a Recombinase Polymerase Amplification Lateral Flow (RPA-LF) assay for the detection of B. divergens . Methods: RPA assays targeting B. divergens hsp70 , cox1 , and 18S rRNA genes were designed and evaluated for sensitivity and specificity using serial dilutions and control samples, including other apicomplexan parasites and tick samples spiked with B. divergens DNA. To assess field performance, 279 I. ricinus ticks were collected from urban parks in Greater London and screened by qPCR and a subset also tested using the new RPA-LF assays. Anaplasma phagocytophilum was screened by qPCR but not detected. Results: The cox1 and 18S rRNA RPA-LF assays demonstrated high specificity, with detection limits of at least 10 - ⁵ ng/μl and 10 - ³ ng/μl, respectively, under isothermal conditions (39 °C for 30 minutes). No cross-reactivity was observed with non-target DNA. Field validation using pooled samples revealed a single sample positive for Babesia divergens or the closely related Babesia capreoli by qPCR, as previously observed in UK tick studies, indicating an estimated nymphal infection prevalence of 0.4% to 1.2%. The 18S rRNA RPA-LF assay successfully detected this pool, whereas the cox1 RPA-LF did not. As with previous molecular approaches, the close genetic similarity of the selected targets prevents discrimination between B. divergens and B. capreoli . Conclusions: We report the first successful development of an RPA-LF assay for B. divergens detection in ticks. The assay demonstrates high sensitivity and specificity under isothermal conditions and provides a portable and reliable tool for field-based molecular surveillance of emerging tick-borne pathogens. Field validation revealed B. divergens / B. capreoli in questing ticks from urban green spaces in London, underscoring the importance of this tool for early detection and ongoing surveillance of tick-borne parasites in expanding urban ecosystems. Babesia divergens RPA-LF Recombinase polymerase amplification Babesiosis Lateral flow Urban Green space Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Background Transmitting a wider variety of pathogens than any other arthropod vector, ticks are second only to mosquitoes in vectoring human and veterinary-relevant pathogens[ 1 ]. Ticks can transmit parasites such as Babesia divergens (bovine and human babesiosis), bacteria such as Borrelia burgdorferi sensu lato (Lyme disease) and Anaplasma phagocytophilum (human granulocytic anaplasmosis and pasture fever), and viral pathogens such as tick-borne encephalitis virus (TBEV). In addition, tick bites can trigger severe allergic reactions, such as alpha-gal syndrome [ 2 ]. In recent decades, expanding ranges of ticks [ 3 ], environmental changes [ 4 ], and the increasing prevalence of coinfection with multiple pathogens within tick populations [ 5 ] have fuelled rises in both the distribution and density of tick-borne diseases (TBD) [ 6 ], raising significant public health concerns and fuelling the initiation of various tick surveillance schemes. Despite this growing threat of TBDs in the face of increased tick/human contact, European surveillance efforts largely focus on screening for Borrelia burgdorferi , as Lyme disease is the most reported and economically significant TBD in the northern hemisphere [ 7 – 9 ]. These efforts have inevitably overshadowed and overlooked other clinically important pathogens such as Babesia divergens .[ 10 ] Historically associated with bovine babesiosis throughout cattle-grazing areas of Europe, Babesia parasites are intraerythrocytic piroplasms that are now recognised as severe human pathogens, potentially life-threatening in immunocompromised individuals[ 11 , 12 ]. While only 60 confirmed cases of human babesiosis have been reported in Europe, and in UK a recent case in Devon [ 10 , 13 ], the asymptomatic seropositivity in patients suggests significant underdiagnosis and underreporting [ 10 , 11 ]. This is particularly important when considering the potential severity in patients with Borrelia burgdorferi coinfections [ 5 ] as high tick exposure individuals are more likely seropositive for multiple TBDs simultaneously [ 5 ]. Once established in tick vectors, Babesia parasites can persist through transstadial and transovarial transmission, allowing infected ticks to maintain the pathogen without requiring reinfection from vertebrate hosts during their development, thereby enhancing transmission potential [ 11 ]. In the UK and Europe, B. divergens transmission is primarily facilitated by Ixodes ricinus . Capable of transmitting B. microti, B. venatorum and B. capreoli [ 10 , 14 ], its extensive range spans from Scandinavia to North Africa [ 2 , 5 ], making it the most widespread and epidemiologically significant tick species in Europe for babesiosis transmission. Furthermore, localised outbreaks of autochthonous canine babesiosis in Essex, South Wales and Devon[ 14 , 15 ], transmitted by Dermacentor reticulatus , underline the need for xenosurveilance to monitor emerging TBDs. Existing systems used to screen ticks for pathogens are primarily PCR-based, which can be inaccessible without specialist infrastructure and technical expertise[ 2 , 16 ]. Whilst such systems have their place as part of established surveillance programmes, the development of a rapid, low-resource, field-deployable molecular assay would provide an additional tool for current surveillance efforts[ 9 , 17 , 18 ]. Such complementary diagnostic approaches would enhance surveillance capabilities by enabling preliminary on-site screening, particularly in resource-limited settings or remote locations where conventional PCR workflows are not feasible, while maintaining the option for confirmatory testing using established laboratory-based methods. Recombinase polymerase amplification (RPA) presents an alternative to PCR for application in field settings, as it requires minimal technical investment due to its rapid DNA amplification (30 min) at constant low temperatures (~ 39°C), thereby eliminating the need for expensive thermocyclers[ 19 ]. Other isothermal methods pose their own unique obstacles, such as loop-mediated isothermal amplification (LAMP), which requires high temperatures (65˚C), complex primer design, and frequent, nonspecific amplification [ 16 ], while Helicase-dependent amplification (HDA) requires complex buffer optimisation [ 20 ]. Thus, RPA is the most promising point-of-need, isothermal molecular assay suited for rapid pathogen surveillance as it operates quickly at a temperature range convenient for field situations [ 16 , 20 ]. Field-portable RPA suitcases, containing the laboratory reagents and equipment needed, are also available to support the deployment of RPA at the point of need, expanding its capabilities to include resource-poor areas[ 16 ]. RPA has previously been used to detect Babesia in agriculture, human and veterinary medicine [ 19 ], including Babesia vogeli, B. gibsoni, B. canis [ 21 , 22 ], B. orientalis [ 23 ] and B. microti [ 24 ]. Coupling RPA with lateral flow (LF) detection systems further enhances its utility as LF strips allow for rapid, equipment-free result interpretation, with gold nanoparticle-based probes providing signal amplification. In fact, LF-RPA assays targeting B. vogeli and B. gibsoni have demonstrated up to 10 5 -fold greater sensitivity than RPA products visualised via gel electrophoresis[ 21 ]. However, to date, no RPA assay has been developed for the detection of Babesia divergens . Given the expansion of tick populations and the importance of Babesia divergens as a zoonotic pathogen, alongside the continued absence of field-portable molecular diagnostic tools, rapid isothermal detection assays would be critical for improving B. divergens surveillance methods. In this study, we established a novel Babesia divergens -specific RPA-LF assay offering a resource-limited alternative to PCR, specifically in the role of xenomonintoring using the presence of pathogen-positive ticks as a proxy for human/animal infection. Previous studies have detected Babesia -positive ticks via PCR in parts of Scotland, Wales, and England, with reported prevalence ranging from 0.2% to 0.38% (2014–2020)[ 10 , 25 , 26 ]. As a proof-of-concept to validate the assay under realistic field conditions, we screened ticks collected from two urban green spaces across Greater London, demonstrating the assay's potential application in routine surveillance and public health monitoring. Methods Tick collection and morphological identification A total of 279 ticks (9 Ixodes ricinus adults, 249 Ixodes ricinus nymphs and 21 Ixodes nymphs) were successfully collected in June 2024 during the first seasonal peak in tick activity in summer [ 6 , 9 , 10 , 27 ]. Considering the recognised risk of urban green spaces as known tick habitats in the UK [ 27 , 28 ], Bushy Park and Richmond Park were selected (see Supplemental Information S1 for coordinates), owing to their substantial deer population, past records of ticks, ease of access and diversity of potential tick habitats [ 1 ]. Questing ticks were collected using the flagging sampling method [ 29 ]. Nymphs were immediately transferred to tubes containing 70% ethanol [ 29 ]. Collected ticks were identified up to the life stage and species level using the identification keys, using a Leitz Laborlux S binocular microscope [ 3 , 30 , 31 ]. Ticks whose species could not be confidently determined were excluded from further study. Due to the challenges associated with identifying tick larvae [ 3 ] [ 31 ] and time constraints, the larvae were discarded. DNA extraction and quantification DNA was extracted from ticks using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, Germany). After storage, transport and identification, field samples were transferred to 100 µl of MilliQ Water (Millipore MilliQ system) in pools of 3 Ixodes ricinus nymphs. Small pool sizes maximised accurate minimum infection rates, pool prevalence and positivity estimates while balancing prevalence underestimation against pool dilution effects [ 32 ]. A total of 200 µl of ATL buffer was added, and samples were homogenised via crushing with pipette tips. Proteinase K (20 µl) was added, and each tube was incubated overnight at 56°C to maximise lysis [ 33 ]. The subsequent steps were followed as per the manufacturer's instructions, except the elution of DNA was performed by adding 30 µl of buffer EB to concentrate DNA. After extraction, DNA was quantified using a Qubit DS-11 FX+ Spectrophotometer/fluorometer (Denovix, USA) using the HS assay. DNA was also extracted from B. divergens (Rouen 1987) grown in vitro in human red blood cells (RBCs). RBCs were supplied by the NHS Blood and Transplant service as research red cells (approval by the research ethics committee, NHS HRA 21/YH/0085 and RVC URN 2021 20044-3). B. divergens infected RBCs at ~ 15% parasitaemia were lysed using 0.075% (w/v) saponin in PBS at room temperature to release haemoglobin, washed in PBS and total genomic DNA was extracted using Qiagen’s QIAamp DNA blood kit as per manufacturer’s instructions. Quantitative PCR DNA controls for Anaplasma phagocytophilum and Babesia divergens were obtained from the Tick Cell Biobank, University of Liverpool. Dilutions were performed for these samples to determine the limit of detections. qPCR analysis of the serial dilution and tick samples were assessed using the Quantinova probe kit (Qiagen) according to the manufacturer’s instructions [ 34 ] with primer/probe combinations as described [ 35 , 36 ] (Supplemental Table S2). Conventional PCR To confirm any qPCR positives, PCR was conducted to test the Babesia genus primers targeting 18S rRNA and Bath-F (Supplemental Table S2) [ 36 , 37 ]. PCR amplification was carried out in a total volume of 25 µl, containing 1.5 µl of sample DNA, 5 µl of Q5 buffer, 0.5 µl of dNTPs, 16.5 µl of water, 0.25 µl of Q5 High Fidelity DNA Polymerase (New England Biolabs), 0.125 µl of forwards primer and 0.125 µl of reverse primer (100µM). Amplification was performed with the following conditions: hold cycle of 90°C for 30s followed by 35 repeats of 98°C for 10s, 54°C for 30s and 72°C for 55s and held at 72°C for 2 mins, on a Kyratec SC300G-R2 PCR thermocycler. Results were analysed in a 1% agarose gel in Tris-acetate-EDTA (TAE). Gels were stained using SYBR Safe DNA Gel Stain (Thermofisher), while Hyperladder 1kb and Hyperladder 100bp (Bioline) were used for size determination. Gels were routinely run at 120 V for 30 minutes and visualised using a Gel Doc XR + Gel Documentation System (BIO-RAD) and Imagelab. RPA primer design Sequences of Babesia divergens HSP70 (EU185809.1), cox1 (MG344907.1), and 18S rRNA (LC477139.1) genes available from Genbank NCBI were aligned using the alignment editor Aliview (Windows version 1.28) and potential primers generated via Primer3. Primers (Supplemental Table 2) were based on the parameters for optimal RPA primers in the TwistAMP design manual [ 38 ]: a GC content between 70%-30%, a primer T m between 50–100°C and no more than 5 mononucleotide repeats, and were synthesised by IDT (Integrated DNA Technologies). Primer stocks were diluted to 100 µM with Milli-Q water and further diluted to 10 µM prior to their use. Primer screening Primer screening (Supplemental table S3) using RPA Basic liquid kit (TwistDx) was performed according to the manufacturer’s instruction [ 39 ] with the single modification that 2.5 µl of each primer was added to the primer mix, and the reaction was run uninterrupted at 39°C for 30 mins in a heat block.. To enable visualisation of amplified product with gel electrophoresis, an additional cycle of 20 mins at 99°C was conducted to denature the proteins and halt the reaction. Probe Design Lateral flow (LF) probes were designed to sit between the forward and reverse primers without overlapping. The probe was designed as advised by TwistDx [ 38 ] where a sequence of 46–52 nucleotides long was chosen, and a tetrahydrofuran (THF) residue was inserted at least 30 bp from the 5’ end and 15 bp from the 3’ end. The 5′ end of the probe was labelled with 5-carboxyfluorescein (FAM), while a C3-spacer was added at the 3′ end. Additionally, the primer on the strand opposite the probe was modified with a biotin label at the 5′ end. The sequence and primers were imported into the online Multiple Primer Analyser [ 40 ] and sequences were repeatedly tested for self-dimerisation and cross-dimerisation. Potential probe sequences were also chosen to lie between as many primer pairs as possible to maximise success, and were analysed via BLAST search for any cross reactions over 95% identity. These sequences (Supplementary table S4) were synthesised by IDT. RPA-LF Most RPA-LF reactions were conducted via modifications to RPA Basic Solid and RPA Basic Liquid protocols. When using the TwistDx RPA Basic solid kit, using 100 µM stocks, a primer-probe master mix consisting 4 µl of forward and biotin primer each, 2 µl of probe and 90 µl of nuclease-free water was made per sample and stored at -20°C. For a 10x RPA Enzyme master mix, 295 µl of TwistDx 2x reaction buffer was added to 30 µl of endonuclease IV (New England Biolabs). A total of 40.5 µl of enzyme master mix was added to each TwistAMP Basic lyophilised pellet tube alongside 5 µl of primer-probe master mix and 2 µl of the DNA sample (~ 2-10ng/ul). When using the TwistDx RPA liquid kit, 100 µM primers and probe stocks were diluted to 10 µM. Per reaction, 21.5 µl of master mix containing 2.4 µl of water, 3 µl of Endonuclease IV, 0.6 µl of diluted probe, 3 µl of dNTPs, 2.5 µl of core reaction mixture, 5 µl of E-MIX, 2.5 µl of diluted forward primer and 2.5 µl of diluted biotinylated reverse primer was added to 1 µl of sample DNA. Using either kit, the reaction is initiated by adding 2.5 µl of magnesium acetate and running at 39°C for 45 mins. After incubation, a PCRD Lateral flow cassette and PCRD FLEX LF dipstick (Abingdon Health, UK) were used to detect RPA products in accordance with the manufacturer's instructions [ 41 ]. These tests have 2 test lines and 1 control line and will detect amplicons that are labelled with DIG/Biotin and/or FITC (or FAM)/Biotin. Sensitivity and Specificity To evaluate the sensitivity of the RPA Assay, 2 µl of Babesia divergens control DNA was added to 18 µl of nuclease-free water in 10x serial dilutions, resulting in concentrations between 10 1 and 10 − 7 ng/µl. Probe specificity was investigated experimentally using genomic DNA extracted from samples of ticks Dermacentor reticulatus, Ixodes ricinus , and from in vitro cultures of laboratory strains of Plasmodium falciparum (3D7 laboratory strain), Plasmodium knowlesi (A1.H1), and B. divergens (Rouen 1987). Results Primer Testing and Probe Selection for B. divergens Detection Thirteen pairs of RPA primers targeting Heat Shock Protein 70 ( hsp70 ), mitochondrial cytochrome-c oxidase subunit 1 gene ( cox1 ), and ribosomal 18S rRNA ( 18S rRNA ) were tested to identify the optimal primer sets for B. divergens detection through RPA assays (Supplementary Table S3). The primers targeting hsp70 failed to produce the expected banding pattern in the presence of the positive control and were therefore unsuitable for further use (Fig. 1 C). For the cox1 target, primer sets Cox1-1, Cox1-3, and Cox1-4 successfully produced bands of the predicted sizes (233, 246 and 249 bp, respectively) (Fig. 1 A, B). Since Cox1-1 and Cox1-3 primers amplified overlapping regions, a probe was designed that could be used with both sets (Supplementary Table S4). Among the 18S rRNA primer pairs, sets 4, 6, and 9 generated the expected bands of 216, 195 and 174 bp, respectively (Fig. 1 B, D). Probes were designed for 18S rRNA-4 (based on Genbank number EF458223.1) and 18S rRNA-6 (based on Genbank number LC477139.1) (Supplementary Table S4). Before testing the cox1 and 18S rRNA probes, BLASTn searches were conducted to check for potential cross-reactions with other species in silico (Table S4). Cox1-4 assays showed significant cross-reactivity with species both within and outside the Babesia genus, whereas Cox1-1 and Cox1-3 demonstrated minimal cross-reactivity, only reacting with B. capreoli (MG344976.1), due to the high sequence similarity between both species (99.83%). Similarly, the probe and primer set for 18S rRNA-6 were preferred over 18S rRNA-4, as set 4 shared over 95% identity with several species outside the Babesia genus (Table S4). Comparing the performance of the RPA-LF cassette and the RPA-LF FLEX dipstick To compare the performance of both probes, RPA reactions using Probes 1 and 6 were tested on both the standard PCRD LF cassette and the PCRD FLEX dipstick against the B. divergens positive control (2.7 ng/µl) and the water negative control. The PCRD FLEX dipsticks was tested as it could serve as a more cost-effective and field-portable alternative to the cassette for subsequent lateral flow visualisation. Using the PCRD LF cassette, reactions with both Probe 1 and Probe 6 were able to successfully detect RPA amplification, and a positive line was observed (Fig. 2 A). Conversely, the PCRD dipstick was unable to detect amplification with Probe 6 in the presence of the positive control and produced a faint positive line for Probe 1 (Fig. 2 B). Due to the limited sensitivity observed with the PCRD FLEX dipstick format, further comparative analysis was discontinued. All subsequent experiments were conducted using the standard PCRD LF cassette. RPA-LF Sensitivity and Specificity To assess the sensitivity of the assay, RPA-LF tests targeting the cox1 and 18S rRNA genes were performed using tenfold serial dilutions of B. divergens DNA. Each concentration was tested in duplicate. In the cox1 assay, the limit of detection (LOD) was determined to be 5.8 × 10⁻⁵ ng/µL, as positive lines remained faintly visible at this concentration but failed to produce a result at 5.11 × 10⁻⁵ ng/µL (Fig. 3 A). For the 18S rRNA assay, initial tests showed visible positive lines down to 2.2 × 10⁻³ ng/µL (Fig. 3 B). However, across replicates, consistent and reproducible bands were only observed down to 1.22 × 10⁻² ng/µL. Below this concentration, positive lines were usual faint or absent. This difference in observed LOD can be attributed to possible variations due to individual test cassettes or other experimental variables, including primer binding strength, amplification efficiency and copy number. To evaluate the specificity of the cox1 and 18s rRNA RPA-LF assays, RPA amplification was performed using purified genomic DNA from various hosts and related pathogens. In both tests (Fig. 4 ), there was no cross-reaction with DNA from Dermacentor reticulatus , Ixodes ricinus , Plasmodium falciparum and Plasmodium knowlesi , with only the genomic DNA of Babesia divergens displaying a positive result. Although DNA from other Babesia species was not available for testing, these results demonstrate that the RPA-LF assays specifically detect B. divergens without cross-reacting with other related apicomplexan parasites or tick host species. Detection of Babesia divergens in Field Samples A total of 249 Ixodes ricinus nymphs were collected from the field and organised into 78 pools of three nymphs each for qPCR analysis. A single three-nymph pool tested positive for B. divergens (Cycle threshold 35, equating 3.02 x10 − 5 ng/µl). Due to the pooled sampling approach, the exact number of infected ticks could not be determined. Based on this result, the estimated nymphal infection prevalence (NIP) ranges from 0.4% to 1.2%. The qPCR-positive pool was subsequently confirmed using conventional PCR (Fig. 5 ) and Sanger sequencing of PCR product to confirm Babesia -genus DNA content. BLAST (NCBI) analysis of the Sanger-sequenced PCR product revealed over 96% sequence identity with Babesia divergens and Babesia capreoli , indicating a high degree of similarity to both closely related species [ 42 ]. Given the high sequence similarity, the positive sample cannot be definitively assigned to B. divergens or B. capreoli without further molecular confirmation with different targets, consistent with findings in other UK surveillance studies[ 10 , 26 , 42 ]. All samples tested were negative for Anaplasma phagocytophilum . To assess the performance of RPA-LF assays in field-collected samples, both the Cox1-1 and 18S rRNA-6 assays were used to evaluate ten pools, including the one positive by qPCR. All pools that tested negative by qPCR were correctly identified as negative by both developed RPA assays. However, the Cox1-1 assay failed to detect Babesia DNA using the qPCR-positive pool, resulting in a false negative (Fig. 6 A). In contrast, the 18S rRNA-6 RPA assay detected this sample as positive, albeit with variable band intensity between replicates. The 18S rRNA-6 assay also successfully identified Babesia DNA in two artificially spiked positive controls (Fig. 6 B), making the newly developed 18S rRNA RPA-LF the most reliable field-adaptable diagnostic tool for detecting active B.divergens infections from DNA isolated from whole vector sampling. Discussion Babesia divergens is of significant zoonotic importance, with infections documented in both deer and cattle across Europe, and occasional severe cases in human infections [ 13 ]. Human cases, while rare (> 60 cases in Europe) [ 10 ], include recent infections in Scotland [ 43 ] and Southwest England in 2020 [ 13 ], demonstrates an ongoing transmission risk despite a low prevalence of Babesia spp. in UK Ixodes ricinus populations [ 9 , 10 , 26 ], underscoring the growing need for effective field-adaptable surveillance tools. This study thus developed and validated a novel RPA-LF molecular diagnostic tool that could enhance current surveillance capabilities with field testing on collected ticks to demonstrate its practical relevance. In this study, two RPA-LF assays targeting cox1 and 18S rRNA were designed for the rapid and sensitive detection of Babesia divergens , while hsp70 primers failed to amplify the target using B. divergens DNA and were discontinued. Serial dilution showed that the cox1 assay demonstrated a 100-fold greater sensitivity than that of the 18s rRNA assay with limits of detection of 10 − 5 ng/µl and 10 − 3 ng/µl, respectively. Although not as sensitive as previously reported RPA-LFs for B. vogeli RPA-LF (LOD: 1x10⁻⁷ ng/µl)[ 21 ], the sensitivity achieved was in line with assays for Anaplasma phagocytophilum (1.77x10⁻⁵ ng/µl), Plasmodium falciparum , and Puccinia striiformis (both 10⁻⁵ ng/µl) [ 44 – 46 ]. Specificity testing confirmed that both cox1 and 18S rRNA RPA-LFs accurately detected B. divergens without cross-reactivity with other apicomplexan DNA ( Plasmodium falciparum , P. knowlesi ) or with local tick species DNA ( Ixodes ricinus , Dermacentor reticulatus ). This aligns with prior Babesia RPA-LF studies showing no cross-reaction with other apicomplexan or tick vectors DNA [ 22 – 24 ]. However, further validation is needed testing other Babesia spp. ( B. gibsoni , B. microti , B. bovis , B. orientalis , B. bigemina ) DNA samples and related piroplasms such as Theileria species present in the UK [ 21 – 24 ]. As most emerging tick-borne pathogens are bacterial or viral and therefore lack 18S rRNA regions [ 8 ], cross-reactivity is unlikely. The RPA-LF assay for both cox1 and 18S rRNA were validated on ten samples, including the qPCR positive sample estimated to a concentration of 10 − 5 ng/µl. The 18S rRNA probe was able to successfully detect this sample alongside two further spiked samples demonstrating diagnostic validity with neither false positives nor false negatives. In contrast, the cox1 probe was unable to detect the field positive sample indicating a lower sensitivity that requires further optimisation with experimentally infected samples. The success of the 18s rRNA assay in the detection of field and spiked samples suggests a sensitivity at approximately 3.02 × 10⁻⁵ ng/µL. The results aligns with scientific literature, with the RPA-LF of Babesia microti , B. vogeli , B. gibsoni and B. orientalis yielding equal or superior sensitivity compared to conventional PCR [ 21 – 24 ]. Further improvement should focus on cox1 primer and probe refinement. The 18S rRNA RPA-LF assays demonstrated fast and reliable detection of B. divergens , completing within 45 minutes at 39°C using pre-prepared reagent mixes. This represents a substantial advance over PCR and LAMP, which require thermal cyclers or constant heat sources. With high specificity for B. divergens and no detectable cross-reactivity, this assay represents an important step towards point-of-care molecular surveillance within vector monitoring programs. Its portability and rapid turnaround make it particularly valuable for xenomonitoring in field settings and remote, resource-limited areas where conventional laboratory infrastructure is lacking. A key barrier to field deployment remains the DNA extraction process, as current protocols rely on commercial kits requiring laboratory equipment such as [ 33 ], vortexers and centrifuges, which limit field applicability. However, RPA’s resilience to crude DNA extracts and biological inhibitors that typically compromise PCR performance [ 16 ] offers significant advantages for field deployment capable of operating with nucleic acids from blood, serum, faecal, nasal, and vaginal swabs, plasma, food, plants, animal tissue, milk, stool and urine [ 16 ]. This robustness also makes it compatible with less refined sample preparations, such as in the detection of malaria parasites from Anopheles mosquitoes extracted using a crude DNA extraction protocol [ 18 ]. Adapting similar extraction protocols for tick samples could streamline field diagnostics and allow for more autonomous surveillance efforts. This compatibility with minimally processed blood samples, such as EDTA or heparinised cow blood samples taken in situ and stored refridgerated in the presence of anticoagulants, would lay the groundwork for a future point-of-care, clinical or veterinary diagnostic tool to complement the limitations of existing serology and microscopy -based diagnostics [ 11 , 12 ]. Finally, tick pooling complicates prevalence estimates. Since ticks were grouped in threes, a single positive pool could reflect one to three infected ticks, implying a nymphal infection prevalence between 0.4% and 1.2%. This range is consistent with previous reports in southern England (0.3–0.46%) and falls within European estimates (0–2.3%) [ 10 , 26 ]. While further sampling is necessary to confidently ascertain the true prevalence of B. divergens in the areas studied, the successful detection of B. divergens in a pooled nymph sample highlights the capability of RPA-LF for monitoring low-prevalence pathogens in field conditions. Conclusion This study successfully developed a Babesia divergens RPA-LF assay that provides a simple, rapid, point-of-sampling tool without the equipment and expertise constraints of PCR. By producing easily interpretable bands in under 45 minutes at 39°C, this project represents a significant step towards a portable molecular diagnostic tool capable of supporting xenomonintoring efforts in diverse surveillance settings. Field validation demonstrated the assay's practical utility by detecting Babesia divergens or B. capreoli in a tick sample from an urban green space, eventually providing future researchers and public health practitioners with a valuable tool for rapid pathogen screening in resource-limited or remote settings where traditional laboratory infrastructure is unavailable. Declarations Ethics approval and consent to participate Red blood cells were supplied by the NHS Blood and Transplant service as research red cells (approval by the research ethics committee, NHS HRA 21/YH/0085 and RVC URN 2021 20044-3). Consent for publication Not Applicable Competing interests The authors declare that they have no competing interests. Funding T.G.C. and S.C. were funded by UKRI grants (ref. BB/X018156/1, MR/X005895/1, MRC IAA2143). EK was supported by the Francis Crick Institute which receives its core funding from Cancer Research UK (FC001003), the UK Medical Research Council (FC001003, and the Wellcome Trust (FC001003). Author Contribution Study conceptualisation:SC,MK. Study methodology:ECJB, SC. Sample collection and species identification: ECJB, KH, CW. Undertaking Experiments: ECJB, SC, EK. Data analysis and curation: ECJB,SC. Writing original draft: ECJB. Project discussion, writing and editing: ECJB,SC,MK, KH,MW, JM, EK, TGC. Supervision: SC, MK, TC. Project administration: SC. Funding Acquisition: SC., TGC. All authors reviewed and approved the final manuscript. Acknowledgement We thank the faculty of infectious and tropical diseases for supporting all facilities and human resources during this study. We are grateful to Elsa Bivas, Alexandra Lynn Hoeger, Luke Brandner Garrod for assisting with the tick sampling for this study. We would also like to thank the NHSBT for the supply of research red cells and the Tick Cell Biobank (University of Liverpool) for supplying DNA controls for Anaplasma phagocytophilum and Babesia divergens. Data Availability All data generated or analysed in this study are included in this published article and supplementary information file. References Greenfield BPJ. Environmental parameters affecting tick (Ixodes ricinus) distribution during the summer season in Richmond Park, London. Bioscience Horizons . 2011;4:140–8. https://doi.org/10.1093/biohorizons/hzr016 Johnson N, Phipps LP, Hansford KM, Folly AJ, Fooks AR, Medlock JM, et al. One Health Approach to Tick and Tick-Borne Disease Surveillance in the United Kingdom. IJERPH . 2022;19:5833. https://doi.org/10.3390/ijerph19105833 Agustín Estrada-Peña, Andrei Daniel Mihalca, Trevor N. Petney. Ticks of Europe and North Africa. New York, NY: Springer Berlin Heidelberg; 2017. Worton AJ, Norman RA, Gilbert L, Porter RB. 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Redescription of Babesia capreoli (Enigk and Friedhoff, 1962) from roe deer (Capreolus capreolus): Isolation, cultivation, host specificity, molecular characterisation and differentiation from Babesia divergens. International Journal for Parasitology. 2010;40:277–84. https://doi.org/10.1016/j.ijpara.2009.08.008 Entrican JH, Williams H, Cook IA, Lancaster WM, Clark JC, Joyner LP, et al. Babesiosis in man: a case from Scotland. BMJ. 1979;2:474–474. https://doi.org/10.1136/bmj.2.6188.474 Kersting S, Rausch V, Bier FF, Von Nickisch-Rosenegk M. Rapid detection of Plasmodium falciparum with isothermal recombinase polymerase amplification and lateral flow analysis. Malar J. 2014;13:99. https://doi.org/10.1186/1475-2875-13-99 Liu Y, Hao J, Guo Q, Yan J, Yao Q. Establishment of a recombinase polymerase amplification detection method for Puccinia striiformis f. sp. tritici. Sci Rep . 2023;13:16133. https://doi.org/10.1038/s41598-023-42663-4 Zhao S, Cui Y, Jing J, Yan Y, Peng Y, Shi K, et al. Rapid and sensitive detection of Anaplasma phagocytophilum using a newly developed recombinase polymerase amplification assay. Experimental Parasitology. 2019;201:21–5. https://doi.org/10.1016/j.exppara.2019.04.010 Additional Declarations No competing interests reported. Supplementary Files 4JanParasitesandvectorsSupplementaryfigures.docx Cite Share Download PDF Status: Under Revision Version 1 posted Editorial decision: Revision requested 19 Apr, 2026 Reviews received at journal 17 Apr, 2026 Reviewers agreed at journal 07 Apr, 2026 Reviews received at journal 07 Apr, 2026 Reviewers agreed at journal 09 Mar, 2026 Reviewers invited by journal 06 Mar, 2026 Editor assigned by journal 02 Mar, 2026 Submission checks completed at journal 02 Mar, 2026 First submitted to journal 01 Mar, 2026 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-8999595","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":604558893,"identity":"3d789b46-182d-4110-b283-9bb2cb660335","order_by":0,"name":"Ernest Chin Jia Bin","email":"","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":false,"prefix":"","firstName":"Ernest","middleName":"Chin Jia","lastName":"Bin","suffix":""},{"id":604558894,"identity":"b6e3d2e7-88d0-40b5-a833-247d5bacf1a9","order_by":1,"name":"Keir Hughes","email":"","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":false,"prefix":"","firstName":"Keir","middleName":"","lastName":"Hughes","suffix":""},{"id":604558896,"identity":"446ca5af-d43d-4586-b96e-3a1de66a10c6","order_by":2,"name":"Cheryl Whitehorn","email":"","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":false,"prefix":"","firstName":"Cheryl","middleName":"","lastName":"Whitehorn","suffix":""},{"id":604558897,"identity":"90b04dc8-7ccb-4622-8119-c7fcd35fab41","order_by":3,"name":"Ellen knuepfer","email":"","orcid":"","institution":"Royal Veterinary College","correspondingAuthor":false,"prefix":"","firstName":"Ellen","middleName":"","lastName":"knuepfer","suffix":""},{"id":604558900,"identity":"69727621-e1a6-489e-8e6e-a92494a58a2a","order_by":4,"name":"Mia White","email":"","orcid":"","institution":"UK Health Security Agency","correspondingAuthor":false,"prefix":"","firstName":"Mia","middleName":"","lastName":"White","suffix":""},{"id":604558901,"identity":"51bc1077-a1d6-400f-8274-4005f611be0d","order_by":5,"name":"Kayleigh Hansford","email":"","orcid":"","institution":"UK Health Security Agency","correspondingAuthor":false,"prefix":"","firstName":"Kayleigh","middleName":"","lastName":"Hansford","suffix":""},{"id":604558902,"identity":"630a5f03-2079-4163-9764-b43c106090ab","order_by":6,"name":"Jolyon Medlock","email":"","orcid":"","institution":"UK Health Security Agency","correspondingAuthor":false,"prefix":"","firstName":"Jolyon","middleName":"","lastName":"Medlock","suffix":""},{"id":604558903,"identity":"a03d3351-c3ff-4a51-81f6-b1d8fdfb1581","order_by":7,"name":"Taane Clark","email":"","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":false,"prefix":"","firstName":"Taane","middleName":"","lastName":"Clark","suffix":""},{"id":604558904,"identity":"6cacf7b5-ea4c-4887-9498-e67e58d19ee4","order_by":8,"name":"Mojca Kristan","email":"","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":false,"prefix":"","firstName":"Mojca","middleName":"","lastName":"Kristan","suffix":""},{"id":604558905,"identity":"35e06fef-3494-4620-93b8-840a0a5f08f1","order_by":9,"name":"Susana campino","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAABDUlEQVRIie3RsUrDQBzH8Z8Imf7RNRLJvcIVIYslz3LlwCxROnbo0CludZMDX0Lf4OSwU2rXg7hIwclBKHQRiolFaCXRjoL33f5wH+6OP+By/cHYaGPQQPcYwddIzYTrbXJGCPZ/Id9mswMJ0udFfwh2eD29N/3BjNjN1cMthgl4oRsJU9lJqCboqPJCGFWUxJ+MZzGR4NNR8zU2Q0geBMqMGz+vSCAr4mnwWbOATefvtIJga/JITNVk9RMRcejnEHxNNMFWZC/XrQ/jxWt86o+Dzl39FyokcStj2xtLOmr7/mU6L2nZZVF5bhY0SCKmei/2bZlEB4Voedln9cY31yBat7LVLmdcLpfrX/YB9mFdjnDLeb8AAAAASUVORK5CYII=","orcid":"","institution":"London School of Hygiene \u0026 Tropical Medicine","correspondingAuthor":true,"prefix":"","firstName":"Susana","middleName":"","lastName":"campino","suffix":""}],"badges":[],"createdAt":"2026-03-01 06:38:24","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-8999595/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-8999595/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":104482762,"identity":"7ace8eff-7763-43f6-9896-075c8ae474d3","added_by":"auto","created_at":"2026-03-12 09:46:23","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":248908,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eAgarose gel electrophoresis screening of RPA primers for \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eB. divergens\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e detection.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(A) COX1 primer sets showing successful amplification of expected amplicon sizes: COX1-1 (233 bp) and COX1-4 (249 bp). (B) Successful amplification of COX1-3 (246 bp) with faint amplification of 18S rRNA-9 (174 bp). (C) Failure of HSP70 primers 1 and 4 to amplify target sequences. (D) 18S rRNA primer evaluation with successful amplification by primer sets 4 (216 bp) and 6 (195 bp). Target genes are labelled above each gel panel with primer pair numbers indicated above individual wells. M = molecular weight marker (sizes indicated in bp); NC- no-template control. Ran on \u003cem\u003eB. divergens\u003c/em\u003e DNA.\u003c/p\u003e","description":"","filename":"image1.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/4074c5a8dda5096043e3fd42.png"},{"id":104781007,"identity":"3cfeaeba-de92-4d32-a079-1d99fceea64f","added_by":"auto","created_at":"2026-03-17 07:54:26","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":232910,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eComparative performance of RPA-LF detection platforms.\u003c/strong\u003e RPA products detected using COX1-Probe 1 and 18S rRNA-Probe 6 with (A) standard PCRD lateral flow cassette and (B) PCRD FLEX dipstick. Positive results show visible test lines; negative controls show only control lines. H₂O = no-template control. Concentrations shown in ng/μL\u003c/p\u003e","description":"","filename":"image2.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/acd60368f113fdece9e1b508.png"},{"id":104482763,"identity":"934b7ad9-b125-4d37-a351-105c84f223bf","added_by":"auto","created_at":"2026-03-12 09:46:23","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":523800,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eLimit of detection determination for \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eB. divergens\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e RPA-LF assays.\u003c/strong\u003e Serial ten-fold dilutions of \u003cem\u003eB. divergens\u003c/em\u003e genomic DNA tested in duplicate using (A) COX1-Probe 1 and (B) 18S rRNA-Probe 6. Concentrations shown in ng/μL. H₂O = negative control.\u003c/p\u003e","description":"","filename":"image3.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/daef1f0580561e3254b15e45.png"},{"id":104482765,"identity":"2002b79b-1081-4c2e-b7a2-696709b3ab2e","added_by":"auto","created_at":"2026-03-12 09:46:23","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":615477,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSpecificity evaluation of \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eB. divergens\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e RPA-LF assays.\u003c/strong\u003eCross-reactivity testing using (A) COX1-Probe 1 and (B) 18S rRNA-Probe 6 against genomic DNA from potential cross-reacting species. Both assays showed specific detection of \u003cem\u003eB. divergens\u003c/em\u003e without cross-reaction to tick species (\u003cem\u003eD. reticulatus, I. ricinus\u003c/em\u003e) or related apicomplexan parasites (\u003cem\u003eP. falciparum,\u003c/em\u003e\u003cdel\u003e\u003cem\u003e \u003c/em\u003e\u003c/del\u003e\u003cem\u003eP. knowlesi\u003c/em\u003e). (+) = \u003cem\u003eB. divergens\u003c/em\u003e positive control; H₂O = negative control.\u003c/p\u003e","description":"","filename":"image4.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/730154b1aaeafeaad0efe618.png"},{"id":104780832,"identity":"f226be7c-4a52-4dd9-b74d-385fe3a40fed","added_by":"auto","created_at":"2026-03-17 07:54:04","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":343975,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003ePCR confirmation of qPCR-positive sample (T31).\u003c/strong\u003e Gel electrophoresis showing amplification products using BATH-F Babesia genus primers (expected size: ~400 bp). Primer targets are labelled above each gel panel M = molecular weight marker; (+) = positive control; NC = negative control; T31 = qPCR positive field sample\u003c/p\u003e","description":"","filename":"image5.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/868d71e2c52af4214ec1bf3c.png"},{"id":104482767,"identity":"3c3de5d6-9e3d-4cb0-b680-00b112fc5a01","added_by":"auto","created_at":"2026-03-12 09:46:23","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":663381,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eRPA-LF validation using field-collected tick samples.\u003c/strong\u003e Performance comparison of (A) COX1-Probe 1 and (B) 18S rRNA-Probe 6 assays. Sample origins (pools): Richmond Park, Bushy Park. The 18S rRNA assay successfully detected the qPCR-positive sample T31 (pool) twice, while COX1 assay failed to detect this low-concentration sample. APC = artificially spiked positive control; (+) = positive controls; H₂O = negative control.\u003c/p\u003e","description":"","filename":"image6.png","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/df7bea3441d28d6845c05cee.png"},{"id":105033019,"identity":"c99e5d39-04d9-4698-bc28-a4578b765f99","added_by":"auto","created_at":"2026-03-20 07:11:38","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":3981343,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/b77563df-5a5d-4b56-8065-bda341a197ae.pdf"},{"id":104482769,"identity":"364f72d4-5e79-4364-9572-ccde89ef8152","added_by":"auto","created_at":"2026-03-12 09:46:23","extension":"docx","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":29935,"visible":true,"origin":"","legend":"","description":"","filename":"4JanParasitesandvectorsSupplementaryfigures.docx","url":"https://assets-eu.researchsquare.com/files/rs-8999595/v1/4d65d9b187e1b9e55c92d064.docx"}],"financialInterests":"No competing interests reported.","formattedTitle":"A Recombinase Polymerase Amplification Lateral Flow Assay (RPA-LF) for detection of Babesia divergens","fulltext":[{"header":"Background","content":"\u003cp\u003eTransmitting a wider variety of pathogens than any other arthropod vector, ticks are second only to mosquitoes in vectoring human and veterinary-relevant pathogens[\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. Ticks can transmit parasites such as \u003cem\u003eBabesia divergens\u003c/em\u003e (bovine and human babesiosis), bacteria such as \u003cem\u003eBorrelia burgdorferi\u003c/em\u003e sensu lato (Lyme disease) and \u003cem\u003eAnaplasma phagocytophilum\u003c/em\u003e (human granulocytic anaplasmosis and pasture fever), and viral pathogens such as tick-borne encephalitis virus (TBEV). In addition, tick bites can trigger severe allergic reactions, such as alpha-gal syndrome [\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eIn recent decades, expanding ranges of ticks [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e], environmental changes [\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e], and the increasing prevalence of coinfection with multiple pathogens within tick populations [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e] have fuelled rises in both the distribution and density of tick-borne diseases (TBD) [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e], raising significant public health concerns and fuelling the initiation of various tick surveillance schemes. Despite this growing threat of TBDs in the face of increased tick/human contact, European surveillance efforts largely focus on screening for \u003cem\u003eBorrelia burgdorferi\u003c/em\u003e, as Lyme disease is the most reported and economically significant TBD in the northern hemisphere [\u003cspan additionalcitationids=\"CR8\" citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e]. These efforts have inevitably overshadowed and overlooked other clinically important pathogens such as \u003cem\u003eBabesia divergens\u003c/em\u003e.[\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e]\u003c/p\u003e \u003cp\u003eHistorically associated with bovine babesiosis throughout cattle-grazing areas of Europe, \u003cem\u003eBabesia\u003c/em\u003e parasites are intraerythrocytic piroplasms that are now recognised as severe human pathogens, potentially life-threatening in immunocompromised individuals[\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e]. While only 60 confirmed cases of human babesiosis have been reported in Europe, and in UK a recent case in Devon [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e], the asymptomatic seropositivity in patients suggests significant underdiagnosis and underreporting [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e]. This is particularly important when considering the potential severity in patients with \u003cem\u003eBorrelia burgdorferi\u003c/em\u003e coinfections [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e] as high tick exposure individuals are more likely seropositive for multiple TBDs simultaneously [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. Once established in tick vectors, \u003cem\u003eBabesia\u003c/em\u003e parasites can persist through transstadial and transovarial transmission, allowing infected ticks to maintain the pathogen without requiring reinfection from vertebrate hosts during their development, thereby enhancing transmission potential [\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e]. In the UK and Europe, \u003cem\u003eB. divergens\u003c/em\u003e transmission is primarily facilitated by \u003cem\u003eIxodes ricinus\u003c/em\u003e. Capable of transmitting \u003cem\u003eB. microti, B. venatorum\u003c/em\u003e and \u003cem\u003eB. capreoli\u003c/em\u003e[\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e], its extensive range spans from Scandinavia to North Africa [\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e, \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e], making it the most widespread and epidemiologically significant tick species in Europe for babesiosis transmission. Furthermore, localised outbreaks of autochthonous canine babesiosis in Essex, South Wales and Devon[\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e], transmitted by \u003cem\u003eDermacentor reticulatus\u003c/em\u003e, underline the need for xenosurveilance to monitor emerging TBDs.\u003c/p\u003e \u003cp\u003eExisting systems used to screen ticks for pathogens are primarily PCR-based, which can be inaccessible without specialist infrastructure and technical expertise[\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e, \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]. Whilst such systems have their place as part of established surveillance programmes, the development of a rapid, low-resource, field-deployable molecular assay would provide an additional tool for current surveillance efforts[\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e, \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e]. Such complementary diagnostic approaches would enhance surveillance capabilities by enabling preliminary on-site screening, particularly in resource-limited settings or remote locations where conventional PCR workflows are not feasible, while maintaining the option for confirmatory testing using established laboratory-based methods.\u003c/p\u003e \u003cp\u003eRecombinase polymerase amplification (RPA) presents an alternative to PCR for application in field settings, as it requires minimal technical investment due to its rapid DNA amplification (30 min) at constant low temperatures (~\u0026thinsp;39\u0026deg;C), thereby eliminating the need for expensive thermocyclers[\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e]. Other isothermal methods pose their own unique obstacles, such as loop-mediated isothermal amplification (LAMP), which requires high temperatures (65˚C), complex primer design, and frequent, nonspecific amplification [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e], while Helicase-dependent amplification (HDA) requires complex buffer optimisation [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e]. Thus, RPA is the most promising point-of-need, isothermal molecular assay suited for rapid pathogen surveillance as it operates quickly at a temperature range convenient for field situations [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e, \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e]. Field-portable RPA suitcases, containing the laboratory reagents and equipment needed, are also available to support the deployment of RPA at the point of need, expanding its capabilities to include resource-poor areas[\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eRPA has previously been used to detect \u003cem\u003eBabesia\u003c/em\u003e in agriculture, human and veterinary medicine [\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e], including \u003cem\u003eBabesia vogeli, B. gibsoni, B. canis\u003c/em\u003e [\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e, \u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e], \u003cem\u003eB. orientalis\u003c/em\u003e [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e] and \u003cem\u003eB. microti\u003c/em\u003e [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. Coupling RPA with lateral flow (LF) detection systems further enhances its utility as LF strips allow for rapid, equipment-free result interpretation, with gold nanoparticle-based probes providing signal amplification. In fact, LF-RPA assays targeting \u003cem\u003eB. vogeli\u003c/em\u003e and \u003cem\u003eB. gibsoni\u003c/em\u003e have demonstrated up to 10\u003csup\u003e5\u003c/sup\u003e-fold greater sensitivity than RPA products visualised via gel electrophoresis[\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e]. However, to date, no RPA assay has been developed for the detection of \u003cem\u003eBabesia divergens\u003c/em\u003e.\u003c/p\u003e \u003cp\u003eGiven the expansion of tick populations and the importance of \u003cem\u003eBabesia divergens\u003c/em\u003e as a zoonotic pathogen, alongside the continued absence of field-portable molecular diagnostic tools, rapid isothermal detection assays would be critical for improving \u003cem\u003eB. divergens\u003c/em\u003e surveillance methods. In this study, we established a novel \u003cem\u003eBabesia divergens\u003c/em\u003e-specific RPA-LF assay offering a resource-limited alternative to PCR, specifically in the role of xenomonintoring using the presence of pathogen-positive ticks as a proxy for human/animal infection. Previous studies have detected \u003cem\u003eBabesia\u003c/em\u003e-positive ticks via PCR in parts of Scotland, Wales, and England, with reported prevalence ranging from 0.2% to 0.38% (2014\u0026ndash;2020)[\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. As a proof-of-concept to validate the assay under realistic field conditions, we screened ticks collected from two urban green spaces across Greater London, demonstrating the assay's potential application in routine surveillance and public health monitoring.\u003c/p\u003e"},{"header":"Methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eTick collection and morphological identification\u003c/h2\u003e \u003cp\u003eA total of 279 ticks (9 \u003cem\u003eIxodes ricinus\u003c/em\u003e adults, 249 \u003cem\u003eIxodes ricinus\u003c/em\u003e nymphs and 21 \u003cem\u003eIxodes\u003c/em\u003e nymphs) were successfully collected in June 2024 during the first seasonal peak in tick activity in summer [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e, \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]. Considering the recognised risk of urban green spaces as known tick habitats in the UK [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e, \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e], Bushy Park and Richmond Park were selected (see Supplemental Information S1 for coordinates), owing to their substantial deer population, past records of ticks, ease of access and diversity of potential tick habitats [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. Questing ticks were collected using the flagging sampling method [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e]. Nymphs were immediately transferred to tubes containing 70% ethanol [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e]. Collected ticks were identified up to the life stage and species level using the identification keys, using a Leitz Laborlux S binocular microscope [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e, \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. Ticks whose species could not be confidently determined were excluded from further study. Due to the challenges associated with identifying tick larvae [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e] [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e] and time constraints, the larvae were discarded.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eDNA extraction and quantification\u003c/h3\u003e\n\u003cp\u003eDNA was extracted from ticks using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, Germany). After storage, transport and identification, field samples were transferred to 100 \u0026micro;l of MilliQ Water (Millipore MilliQ system) in pools of 3 \u003cem\u003eIxodes ricinus\u003c/em\u003e nymphs. Small pool sizes maximised accurate minimum infection rates, pool prevalence and positivity estimates while balancing prevalence underestimation against pool dilution effects [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. A total of 200 \u0026micro;l of ATL buffer was added, and samples were homogenised via crushing with pipette tips. Proteinase K (20 \u0026micro;l) was added, and each tube was incubated overnight at 56\u0026deg;C to maximise lysis [\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]. The subsequent steps were followed as per the manufacturer's instructions, except the elution of DNA was performed by adding 30 \u0026micro;l of buffer EB to concentrate DNA. After extraction, DNA was quantified using a Qubit DS-11 FX+ Spectrophotometer/fluorometer (Denovix, USA) using the HS assay. DNA was also extracted from \u003cem\u003eB. divergens\u003c/em\u003e (Rouen 1987) grown \u003cem\u003ein vitro\u003c/em\u003e in human red blood cells (RBCs). RBCs were supplied by the NHS Blood and Transplant service as research red cells (approval by the research ethics committee, NHS HRA 21/YH/0085 and RVC URN 2021 20044-3). \u003cem\u003eB. divergens\u003c/em\u003e infected RBCs at ~\u0026thinsp;15% parasitaemia were lysed using 0.075% (w/v) saponin in PBS at room temperature to release haemoglobin, washed in PBS and total genomic DNA was extracted using Qiagen\u0026rsquo;s QIAamp DNA blood kit as per manufacturer\u0026rsquo;s instructions.\u003c/p\u003e\n\u003ch3\u003eQuantitative PCR\u003c/h3\u003e\n\u003cp\u003eDNA controls for \u003cem\u003eAnaplasma phagocytophilum\u003c/em\u003e and \u003cem\u003eBabesia divergens\u003c/em\u003e were obtained from the Tick Cell Biobank, University of Liverpool. Dilutions were performed for these samples to determine the limit of detections. qPCR analysis of the serial dilution and tick samples were assessed using the Quantinova probe kit (Qiagen) according to the manufacturer\u0026rsquo;s instructions [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e] with primer/probe combinations as described [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e] (Supplemental Table S2).\u003c/p\u003e\n\u003ch3\u003eConventional PCR\u003c/h3\u003e\n\u003cp\u003eTo confirm any qPCR positives, PCR was conducted to test the \u003cem\u003eBabesia\u003c/em\u003e genus primers targeting \u003cem\u003e18S rRNA\u003c/em\u003e and \u003cem\u003eBath-F\u003c/em\u003e (Supplemental Table S2) [\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e, \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e]. PCR amplification was carried out in a total volume of 25 \u0026micro;l, containing 1.5 \u0026micro;l of sample DNA, 5 \u0026micro;l of Q5 buffer, 0.5 \u0026micro;l of dNTPs, 16.5 \u0026micro;l of water, 0.25 \u0026micro;l of Q5 High Fidelity DNA Polymerase (New England Biolabs), 0.125 \u0026micro;l of forwards primer and 0.125 \u0026micro;l of reverse primer (100\u0026micro;M). Amplification was performed with the following conditions: hold cycle of 90\u0026deg;C for 30s followed by 35 repeats of 98\u0026deg;C for 10s, 54\u0026deg;C for 30s and 72\u0026deg;C for 55s and held at 72\u0026deg;C for 2 mins, on a Kyratec SC300G-R2 PCR thermocycler. Results were analysed in a 1% agarose gel in Tris-acetate-EDTA (TAE). Gels were stained using SYBR Safe DNA Gel Stain (Thermofisher), while Hyperladder 1kb and Hyperladder 100bp (Bioline) were used for size determination. Gels were routinely run at 120 V for 30 minutes and visualised using a Gel Doc XR\u0026thinsp;+\u0026thinsp;Gel Documentation System (BIO-RAD) and Imagelab.\u003c/p\u003e\n\u003ch3\u003eRPA primer design\u003c/h3\u003e\n\u003cp\u003eSequences of \u003cem\u003eBabesia divergens HSP70\u003c/em\u003e (EU185809.1), \u003cem\u003ecox1\u003c/em\u003e (MG344907.1), and \u003cem\u003e18S rRNA\u003c/em\u003e (LC477139.1) genes available from Genbank NCBI were aligned using the alignment editor Aliview (Windows version 1.28) and potential primers generated via Primer3. Primers (Supplemental Table\u0026nbsp;2) were based on the parameters for optimal RPA primers in the TwistAMP design manual [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e]: a GC content between 70%-30%, a primer T\u003csub\u003em\u003c/sub\u003e between 50\u0026ndash;100\u0026deg;C and no more than 5 mononucleotide repeats, and were synthesised by IDT (Integrated DNA Technologies). Primer stocks were diluted to 100 \u0026micro;M with Milli-Q water and further diluted to 10 \u0026micro;M prior to their use.\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003ePrimer screening\u003c/h2\u003e \u003cp\u003ePrimer screening (Supplemental table S3) using RPA Basic liquid kit (TwistDx) was performed according to the manufacturer\u0026rsquo;s instruction [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e] with the single modification that 2.5 \u0026micro;l of each primer was added to the primer mix, and the reaction was run uninterrupted at 39\u0026deg;C for 30 mins in a heat block.. To enable visualisation of amplified product with gel electrophoresis, an additional cycle of 20 mins at 99\u0026deg;C was conducted to denature the proteins and halt the reaction.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eProbe Design\u003c/h3\u003e\n\u003cp\u003eLateral flow (LF) probes were designed to sit between the forward and reverse primers without overlapping. The probe was designed as advised by TwistDx [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e] where a sequence of 46\u0026ndash;52 nucleotides long was chosen, and a tetrahydrofuran (THF) residue was inserted at least 30 bp from the 5\u0026rsquo; end and 15 bp from the 3\u0026rsquo; end. The 5\u0026prime; end of the probe was labelled with 5-carboxyfluorescein (FAM), while a C3-spacer was added at the 3\u0026prime; end. Additionally, the primer on the strand opposite the probe was modified with a biotin label at the 5\u0026prime; end. The sequence and primers were imported into the online Multiple Primer Analyser [\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e] and sequences were repeatedly tested for self-dimerisation and cross-dimerisation. Potential probe sequences were also chosen to lie between as many primer pairs as possible to maximise success, and were analysed via BLAST search for any cross reactions over 95% identity. These sequences (Supplementary table S4) were synthesised by IDT.\u003c/p\u003e\n\u003ch3\u003eRPA-LF\u003c/h3\u003e\n\u003cp\u003eMost RPA-LF reactions were conducted via modifications to RPA Basic Solid and RPA Basic Liquid protocols. When using the TwistDx RPA Basic solid kit, using 100 \u0026micro;M stocks, a primer-probe master mix consisting 4 \u0026micro;l of forward and biotin primer each, 2 \u0026micro;l of probe and 90 \u0026micro;l of nuclease-free water was made per sample and stored at -20\u0026deg;C. For a 10x RPA Enzyme master mix, 295 \u0026micro;l of TwistDx 2x reaction buffer was added to 30 \u0026micro;l of endonuclease IV (New England Biolabs). A total of 40.5 \u0026micro;l of enzyme master mix was added to each TwistAMP Basic lyophilised pellet tube alongside 5 \u0026micro;l of primer-probe master mix and 2 \u0026micro;l of the DNA sample (~\u0026thinsp;2-10ng/ul). When using the TwistDx RPA liquid kit, 100 \u0026micro;M primers and probe stocks were diluted to 10 \u0026micro;M. Per reaction, 21.5 \u0026micro;l of master mix containing 2.4 \u0026micro;l of water, 3 \u0026micro;l of Endonuclease IV, 0.6 \u0026micro;l of diluted probe, 3 \u0026micro;l of dNTPs, 2.5 \u0026micro;l of core reaction mixture, 5 \u0026micro;l of E-MIX, 2.5 \u0026micro;l of diluted forward primer and 2.5 \u0026micro;l of diluted biotinylated reverse primer was added to 1 \u0026micro;l of sample DNA. Using either kit, the reaction is initiated by adding 2.5 \u0026micro;l of magnesium acetate and running at 39\u0026deg;C for 45 mins. After incubation, a PCRD Lateral flow cassette and PCRD FLEX LF dipstick (Abingdon Health, UK) were used to detect RPA products in accordance with the manufacturer's instructions [\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]. These tests have 2 test lines and 1 control line and will detect amplicons that are labelled with DIG/Biotin and/or FITC (or FAM)/Biotin.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eSensitivity and Specificity\u003c/h2\u003e \u003cp\u003eTo evaluate the sensitivity of the RPA Assay, 2 \u0026micro;l of \u003cem\u003eBabesia divergens\u003c/em\u003e control DNA was added to 18 \u0026micro;l of nuclease-free water in 10x serial dilutions, resulting in concentrations between 10\u003csup\u003e1\u003c/sup\u003e and 10\u003csup\u003e\u0026minus;\u0026thinsp;7\u003c/sup\u003e ng/\u0026micro;l. Probe specificity was investigated experimentally using genomic DNA extracted from samples of ticks \u003cem\u003eDermacentor reticulatus, Ixodes ricinus\u003c/em\u003e, and from \u003cem\u003ein vitro\u003c/em\u003e cultures of laboratory strains of \u003cem\u003ePlasmodium falciparum\u003c/em\u003e (3D7 laboratory strain), \u003cem\u003ePlasmodium knowlesi\u003c/em\u003e (A1.H1), and \u003cem\u003eB. divergens\u003c/em\u003e (Rouen 1987).\u003c/p\u003e \u003c/div\u003e"},{"header":"Results","content":"\u003cp\u003e \u003cb\u003ePrimer Testing and Probe Selection for\u003c/b\u003e \u003cb\u003eB. divergens\u003c/b\u003e \u003cb\u003eDetection\u003c/b\u003e\u003c/p\u003e \u003cp\u003eThirteen pairs of RPA primers targeting Heat Shock Protein 70 (\u003cem\u003ehsp70\u003c/em\u003e), mitochondrial cytochrome-c oxidase subunit 1 gene (\u003cem\u003ecox1\u003c/em\u003e), and ribosomal 18S rRNA (\u003cem\u003e18S rRNA\u003c/em\u003e) were tested to identify the optimal primer sets for \u003cem\u003eB. divergens\u003c/em\u003e detection through RPA assays (Supplementary Table S3). The primers targeting \u003cem\u003ehsp70\u003c/em\u003e failed to produce the expected banding pattern in the presence of the positive control and were therefore unsuitable for further use (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eC). For the \u003cem\u003ecox1\u003c/em\u003e target, primer sets Cox1-1, Cox1-3, and Cox1-4 successfully produced bands of the predicted sizes (233, 246 and 249 bp, respectively) (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eA, B). Since Cox1-1 and Cox1-3 primers amplified overlapping regions, a probe was designed that could be used with both sets (Supplementary Table S4). Among the \u003cem\u003e18S rRNA\u003c/em\u003e primer pairs, sets 4, 6, and 9 generated the expected bands of 216, 195 and 174 bp, respectively (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e1\u003c/span\u003eB, D). Probes were designed for 18S rRNA-4 (based on Genbank number EF458223.1) and 18S rRNA-6 (based on Genbank number LC477139.1) (Supplementary Table S4). Before testing the \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e probes, BLASTn searches were conducted to check for potential cross-reactions with other species in silico (Table S4). Cox1-4 assays showed significant cross-reactivity with species both within and outside the \u003cem\u003eBabesia\u003c/em\u003e genus, whereas Cox1-1 and Cox1-3 demonstrated minimal cross-reactivity, only reacting with \u003cem\u003eB. capreoli\u003c/em\u003e (MG344976.1), due to the high sequence similarity between both species (99.83%). Similarly, the probe and primer set for 18S rRNA-6 were preferred over 18S rRNA-4, as set 4 shared over 95% identity with several species outside the \u003cem\u003eBabesia\u003c/em\u003e genus (Table S4).\u003c/p\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eComparing the performance of the RPA-LF cassette and the RPA-LF FLEX dipstick\u003c/h2\u003e \u003cp\u003eTo compare the performance of both probes, RPA reactions using Probes 1 and 6 were tested on both the standard PCRD LF cassette and the PCRD FLEX dipstick against the \u003cem\u003eB. divergens\u003c/em\u003e positive control (2.7 ng/\u0026micro;l) and the water negative control. The PCRD FLEX dipsticks was tested as it could serve as a more cost-effective and field-portable alternative to the cassette for subsequent lateral flow visualisation. Using the PCRD LF cassette, reactions with both Probe 1 and Probe 6 were able to successfully detect RPA amplification, and a positive line was observed (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). Conversely, the PCRD dipstick was unable to detect amplification with Probe 6 in the presence of the positive control and produced a faint positive line for Probe 1 (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e2\u003c/span\u003eB). Due to the limited sensitivity observed with the PCRD FLEX dipstick format, further comparative analysis was discontinued. All subsequent experiments were conducted using the standard PCRD LF cassette.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eRPA-LF Sensitivity and Specificity\u003c/h2\u003e \u003cp\u003eTo assess the sensitivity of the assay, RPA-LF tests targeting the \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e genes were performed using tenfold serial dilutions of \u003cem\u003eB. divergens\u003c/em\u003e DNA. Each concentration was tested in duplicate. In the \u003cem\u003ecox1\u003c/em\u003e assay, the limit of detection (LOD) was determined to be 5.8 \u0026times; 10⁻⁵ ng/\u0026micro;L, as positive lines remained faintly visible at this concentration but failed to produce a result at 5.11 \u0026times; 10⁻⁵ ng/\u0026micro;L (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). For the \u003cem\u003e18S rRNA\u003c/em\u003e assay, initial tests showed visible positive lines down to 2.2 \u0026times; 10⁻\u0026sup3; ng/\u0026micro;L (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e3\u003c/span\u003eB). However, across replicates, consistent and reproducible bands were only observed down to 1.22 \u0026times; 10⁻\u0026sup2; ng/\u0026micro;L. Below this concentration, positive lines were usual faint or absent. This difference in observed LOD can be attributed to possible variations due to individual test cassettes or other experimental variables, including primer binding strength, amplification efficiency and copy number.\u003c/p\u003e \u003cp\u003eTo evaluate the specificity of the \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18s rRNA\u003c/em\u003e RPA-LF assays, RPA amplification was performed using purified genomic DNA from various hosts and related pathogens. In both tests (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e4\u003c/span\u003e), there was no cross-reaction with DNA from \u003cem\u003eDermacentor reticulatus\u003c/em\u003e, \u003cem\u003eIxodes ricinus\u003c/em\u003e, \u003cem\u003ePlasmodium falciparum\u003c/em\u003e and \u003cem\u003ePlasmodium knowlesi\u003c/em\u003e, with only the genomic DNA of \u003cem\u003eBabesia divergens\u003c/em\u003e displaying a positive result. Although DNA from other \u003cem\u003eBabesia\u003c/em\u003e species was not available for testing, these results demonstrate that the RPA-LF assays specifically detect \u003cem\u003eB. divergens\u003c/em\u003e without cross-reacting with other related apicomplexan parasites or tick host species.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eDetection of Babesia divergens in Field Samples\u003c/h2\u003e \u003cp\u003eA total of 249 \u003cem\u003eIxodes ricinus\u003c/em\u003e nymphs were collected from the field and organised into 78 pools of three nymphs each for qPCR analysis. A single three-nymph pool tested positive for \u003cem\u003eB. divergens\u003c/em\u003e (Cycle threshold 35, equating 3.02 x10\u003csup\u003e\u0026minus;\u0026thinsp;5\u003c/sup\u003e ng/\u0026micro;l). Due to the pooled sampling approach, the exact number of infected ticks could not be determined. Based on this result, the estimated nymphal infection prevalence (NIP) ranges from 0.4% to 1.2%. The qPCR-positive pool was subsequently confirmed using conventional PCR (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e5\u003c/span\u003e) and Sanger sequencing of PCR product to confirm \u003cem\u003eBabesia\u003c/em\u003e-genus DNA content. BLAST (NCBI) analysis of the Sanger-sequenced PCR product revealed over 96% sequence identity with \u003cem\u003eBabesia divergens\u003c/em\u003e and \u003cem\u003eBabesia capreoli\u003c/em\u003e, indicating a high degree of similarity to both closely related species [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. Given the high sequence similarity, the positive sample cannot be definitively assigned to \u003cem\u003eB. divergens\u003c/em\u003e or \u003cem\u003eB. capreoli\u003c/em\u003e without further molecular confirmation with different targets, consistent with findings in other UK surveillance studies[\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e, \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. All samples tested were negative for \u003cem\u003eAnaplasma phagocytophilum\u003c/em\u003e.\u003c/p\u003e \u003cp\u003eTo assess the performance of RPA-LF assays in field-collected samples, both the Cox1-1 and 18S rRNA-6 assays were used to evaluate ten pools, including the one positive by qPCR. All pools that tested negative by qPCR were correctly identified as negative by both developed RPA assays. However, the Cox1-1 assay failed to detect \u003cem\u003eBabesia\u003c/em\u003e DNA using the qPCR-positive pool, resulting in a false negative (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e6\u003c/span\u003eA). In contrast, the 18S rRNA-6 RPA assay detected this sample as positive, albeit with variable band intensity between replicates. The 18S rRNA-6 assay also successfully identified \u003cem\u003eBabesia\u003c/em\u003e DNA in two artificially spiked positive controls (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e6\u003c/span\u003eB), making the newly developed \u003cem\u003e18S rRNA\u003c/em\u003e RPA-LF the most reliable field-adaptable diagnostic tool for detecting active \u003cem\u003eB.divergens\u003c/em\u003e infections from DNA isolated from whole vector sampling.\u003c/p\u003e \u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003e \u003cem\u003eBabesia divergens\u003c/em\u003e is of significant zoonotic importance, with infections documented in both deer and cattle across Europe, and occasional severe cases in human infections [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e]. Human cases, while rare (\u0026gt;\u0026thinsp;60 cases in Europe) [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e], include recent infections in Scotland [\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e] and Southwest England in 2020 [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e], demonstrates an ongoing transmission risk despite a low prevalence of \u003cem\u003eBabesia\u003c/em\u003e spp. in UK \u003cem\u003eIxodes ricinus\u003c/em\u003e populations [\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e], underscoring the growing need for effective field-adaptable surveillance tools. This study thus developed and validated a novel RPA-LF molecular diagnostic tool that could enhance current surveillance capabilities with field testing on collected ticks to demonstrate its practical relevance.\u003c/p\u003e \u003cp\u003eIn this study, two RPA-LF assays targeting \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e were designed for the rapid and sensitive detection of \u003cem\u003eBabesia divergens\u003c/em\u003e, while \u003cem\u003ehsp70\u003c/em\u003e primers failed to amplify the target using \u003cem\u003eB. divergens\u003c/em\u003e DNA and were discontinued. Serial dilution showed that the \u003cem\u003ecox1\u003c/em\u003e assay demonstrated a 100-fold greater sensitivity than that of the \u003cem\u003e18s rRNA\u003c/em\u003e assay with limits of detection of 10\u003csup\u003e\u0026minus;\u0026thinsp;5\u003c/sup\u003eng/\u0026micro;l and 10\u003csup\u003e\u0026minus;\u0026thinsp;3\u003c/sup\u003eng/\u0026micro;l, respectively. Although not as sensitive as previously reported RPA-LFs for \u003cem\u003eB. vogeli\u003c/em\u003e RPA-LF (LOD: 1x10⁻⁷ ng/\u0026micro;l)[\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e], the sensitivity achieved was in line with assays for \u003cem\u003eAnaplasma phagocytophilum\u003c/em\u003e (1.77x10⁻⁵ ng/\u0026micro;l), \u003cem\u003ePlasmodium falciparum\u003c/em\u003e, and \u003cem\u003ePuccinia striiformis\u003c/em\u003e (both 10⁻⁵ ng/\u0026micro;l) [\u003cspan additionalcitationids=\"CR45\" citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eSpecificity testing confirmed that both \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e RPA-LFs accurately detected \u003cem\u003eB. divergens\u003c/em\u003e without cross-reactivity with other apicomplexan DNA (\u003cem\u003ePlasmodium falciparum\u003c/em\u003e, \u003cem\u003eP. knowlesi\u003c/em\u003e) or with local tick species DNA (\u003cem\u003eIxodes ricinus\u003c/em\u003e, \u003cem\u003eDermacentor reticulatus\u003c/em\u003e). This aligns with prior \u003cem\u003eBabesia\u003c/em\u003e RPA-LF studies showing no cross-reaction with other apicomplexan or tick vectors DNA [\u003cspan additionalcitationids=\"CR23\" citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. However, further validation is needed testing other \u003cem\u003eBabesia\u003c/em\u003e spp. (\u003cem\u003eB. gibsoni\u003c/em\u003e, \u003cem\u003eB. microti\u003c/em\u003e, \u003cem\u003eB. bovis\u003c/em\u003e, \u003cem\u003eB. orientalis\u003c/em\u003e, \u003cem\u003eB. bigemina\u003c/em\u003e) DNA samples and related piroplasms such as \u003cem\u003eTheileria\u003c/em\u003e species present in the UK [\u003cspan additionalcitationids=\"CR22 CR23\" citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. As most emerging tick-borne pathogens are bacterial or viral and therefore lack \u003cem\u003e18S rRNA\u003c/em\u003e regions [\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e], cross-reactivity is unlikely.\u003c/p\u003e \u003cp\u003eThe RPA-LF assay for both \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e were validated on ten samples, including the qPCR positive sample estimated to a concentration of 10\u003csup\u003e\u0026minus;\u0026thinsp;5\u003c/sup\u003eng/\u0026micro;l. The \u003cem\u003e18S rRNA\u003c/em\u003e probe was able to successfully detect this sample alongside two further spiked samples demonstrating diagnostic validity with neither false positives nor false negatives. In contrast, the \u003cem\u003ecox1\u003c/em\u003e probe was unable to detect the field positive sample indicating a lower sensitivity that requires further optimisation with experimentally infected samples. The success of the \u003cem\u003e18s rRNA\u003c/em\u003e assay in the detection of field and spiked samples suggests a sensitivity at approximately 3.02 \u0026times; 10⁻⁵ ng/\u0026micro;L. The results aligns with scientific literature, with the RPA-LF of \u003cem\u003eBabesia microti\u003c/em\u003e, \u003cem\u003eB. vogeli\u003c/em\u003e, \u003cem\u003eB. gibsoni\u003c/em\u003e and \u003cem\u003eB. orientalis\u003c/em\u003e yielding equal or superior sensitivity compared to conventional PCR [\u003cspan additionalcitationids=\"CR22 CR23\" citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. Further improvement should focus on \u003cem\u003ecox1\u003c/em\u003e primer and probe refinement.\u003c/p\u003e \u003cp\u003eThe \u003cem\u003e18S rRNA\u003c/em\u003e RPA-LF assays demonstrated fast and reliable detection of \u003cem\u003eB. divergens\u003c/em\u003e, completing within 45 minutes at 39\u0026deg;C using pre-prepared reagent mixes. This represents a substantial advance over PCR and LAMP, which require thermal cyclers or constant heat sources. With high specificity for \u003cem\u003eB. divergens\u003c/em\u003e and no detectable cross-reactivity, this assay represents an important step towards point-of-care molecular surveillance within vector monitoring programs. Its portability and rapid turnaround make it particularly valuable for xenomonitoring in field settings and remote, resource-limited areas where conventional laboratory infrastructure is lacking.\u003c/p\u003e \u003cp\u003eA key barrier to field deployment remains the DNA extraction process, as current protocols rely on commercial kits requiring laboratory equipment such as [\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e], vortexers and centrifuges, which limit field applicability. However, RPA\u0026rsquo;s resilience to crude DNA extracts and biological inhibitors that typically compromise PCR performance [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e] offers significant advantages for field deployment capable of operating with nucleic acids from blood, serum, faecal, nasal, and vaginal swabs, plasma, food, plants, animal tissue, milk, stool and urine [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]. This robustness also makes it compatible with less refined sample preparations, such as in the detection of malaria parasites from \u003cem\u003eAnopheles\u003c/em\u003e mosquitoes extracted using a crude DNA extraction protocol [\u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e]. Adapting similar extraction protocols for tick samples could streamline field diagnostics and allow for more autonomous surveillance efforts. This compatibility with minimally processed blood samples, such as EDTA or heparinised cow blood samples taken in situ and stored refridgerated in the presence of anticoagulants, would lay the groundwork for a future point-of-care, clinical or veterinary diagnostic tool to complement the limitations of existing serology and microscopy -based diagnostics [\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eFinally, tick pooling complicates prevalence estimates. Since ticks were grouped in threes, a single positive pool could reflect one to three infected ticks, implying a nymphal infection prevalence between 0.4% and 1.2%. This range is consistent with previous reports in southern England (0.3\u0026ndash;0.46%) and falls within European estimates (0\u0026ndash;2.3%) [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. While further sampling is necessary to confidently ascertain the true prevalence of \u003cem\u003eB. divergens\u003c/em\u003e in the areas studied, the successful detection of \u003cem\u003eB. divergens\u003c/em\u003e in a pooled nymph sample highlights the capability of RPA-LF for monitoring low-prevalence pathogens in field conditions.\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eThis study successfully developed a \u003cem\u003eBabesia divergens\u003c/em\u003e RPA-LF assay that provides a simple, rapid, point-of-sampling tool without the equipment and expertise constraints of PCR. By producing easily interpretable bands in under 45 minutes at 39\u0026deg;C, this project represents a significant step towards a portable molecular diagnostic tool capable of supporting xenomonintoring efforts in diverse surveillance settings. Field validation demonstrated the assay's practical utility by detecting \u003cem\u003eBabesia divergens\u003c/em\u003e or \u003cem\u003eB. capreoli\u003c/em\u003e in a tick sample from an urban green space, eventually providing future researchers and public health practitioners with a valuable tool for rapid pathogen screening in resource-limited or remote settings where traditional laboratory infrastructure is unavailable.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRed blood cells were supplied by the NHS Blood and Transplant service as research red cells (approval by the research ethics committee, NHS HRA 21/YH/0085 and RVC URN 2021 20044-3).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot Applicable\u003c/p\u003e\n\u003ch2\u003eCompeting interests\u003c/h2\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003ch2\u003eFunding\u003c/h2\u003e\n\u003cp\u003eT.G.C. and S.C. were funded by UKRI grants (ref. BB/X018156/1, MR/X005895/1, MRC IAA2143). EK was supported by the Francis Crick Institute which receives its core funding from Cancer Research UK (FC001003), the UK Medical Research Council (FC001003, and the Wellcome Trust (FC001003).\u003c/p\u003e\n\u003ch2\u003eAuthor Contribution\u003c/h2\u003e\n\u003cp\u003eStudy conceptualisation:SC,MK. Study methodology:ECJB, SC. Sample collection and species identification: ECJB, KH, CW. Undertaking Experiments: ECJB, SC, EK. Data analysis and curation: ECJB,SC. Writing original draft: ECJB. Project discussion, writing and editing: ECJB,SC,MK, KH,MW, JM, EK, TGC. Supervision: SC, MK, TC. Project administration: SC. Funding Acquisition: SC., TGC. All authors reviewed and approved the final manuscript.\u003c/p\u003e\n\u003ch2\u003eAcknowledgement\u003c/h2\u003e\n\u003cp\u003eWe thank the faculty of infectious and tropical diseases for supporting all facilities and human resources during this study. We are grateful to Elsa Bivas, Alexandra Lynn Hoeger, Luke Brandner Garrod for assisting with the tick sampling for this study. We would also like to thank the NHSBT for the supply of research red cells and the Tick Cell Biobank (University of Liverpool) for supplying DNA controls for Anaplasma phagocytophilum and Babesia divergens.\u003c/p\u003e\n\u003ch2\u003eData Availability\u003c/h2\u003e\n\u003cp\u003eAll data generated or analysed in this study are included in this published article and supplementary information file.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eGreenfield BPJ. 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Rapid and sensitive detection of Anaplasma phagocytophilum using a newly developed recombinase polymerase amplification assay. \u003cem\u003eExperimental Parasitology.\u003c/em\u003e 2019;201:21\u0026ndash;5. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1016/j.exppara.2019.04.010\u003c/span\u003e\u003cspan address=\"10.1016/j.exppara.2019.04.010\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"parasites-and-vectors","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"parv","sideBox":"Learn more about [Parasites \u0026 Vectors](http://parasitesandvectors.biomedcentral.com/)","snPcode":"13071","submissionUrl":"https://submission.nature.com/new-submission/13071/3","title":"Parasites \u0026 Vectors","twitterHandle":"@bugbittentweets","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"Babesia divergens, RPA-LF, Recombinase polymerase amplification, Babesiosis, Lateral flow, Urban Green space","lastPublishedDoi":"10.21203/rs.3.rs-8999595/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-8999595/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003e\u003cstrong\u003eBackground:\u003c/strong\u003e \u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eBabesia divergens\u003c/em\u003e is a tick-borne apicomplexan parasite found predominantly in Europe, causing babesiosis in livestock and occasionally in humans, with severe disease reported in immunosuppressed or asplenic individuals. As populations of \u003cem\u003eIxodes ricinus\u003c/em\u003e, the principal vector, expand geographically including into urban environments, there is a growing need for rapid, sensitive, and field-deployable surveillance tools. In this study, we developed a Recombinase Polymerase Amplification Lateral Flow (RPA-LF) assay for the detection of \u003cem\u003eB. divergens\u003c/em\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMethods:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRPA assays targeting \u003cem\u003eB. divergens\u003c/em\u003e \u003cem\u003ehsp70\u003c/em\u003e, \u003cem\u003ecox1\u003c/em\u003e, and \u003cem\u003e18S rRNA\u003c/em\u003e genes were designed and evaluated for sensitivity and specificity using serial dilutions and control samples, including other apicomplexan\u003cem\u003e \u003c/em\u003eparasites and tick samples spiked with \u003cem\u003eB. divergens\u003c/em\u003e DNA. \u0026nbsp;To assess field performance, 279 \u003cem\u003eI. ricinus\u003c/em\u003e ticks were collected from urban parks in Greater London and screened by qPCR and a subset also tested using the new RPA-LF assays. \u003cem\u003eAnaplasma phagocytophilum\u003c/em\u003e was screened by qPCR but not detected.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eResults:\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe \u003cem\u003ecox1\u003c/em\u003e and \u003cem\u003e18S rRNA\u003c/em\u003e RPA-LF assays demonstrated high specificity, with detection limits of at least 10\u003csup\u003e-\u003c/sup\u003e⁵ ng/μl and 10\u003csup\u003e-\u003c/sup\u003e³ ng/μl, respectively, under isothermal conditions (39 °C for 30 minutes). No cross-reactivity was observed with non-target DNA. Field validation using pooled samples revealed a single sample positive for \u003cem\u003eBabesia divergens\u003c/em\u003e or the closely related \u003cem\u003eBabesia capreoli\u003c/em\u003e by qPCR, as previously observed in UK tick studies, indicating an estimated nymphal infection prevalence of 0.4% to 1.2%. The \u003cem\u003e18S rRNA\u003c/em\u003e RPA-LF assay successfully detected this pool, \u0026nbsp;whereas the \u003cem\u003ecox1\u003c/em\u003e RPA-LF did not. As with previous molecular approaches, the close genetic similarity of the selected targets prevents discrimination between \u003cem\u003eB. divergens\u003c/em\u003e and \u003cem\u003eB. capreoli\u003c/em\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConclusions:\u003c/strong\u003e\u003cbr\u003e\nWe report the first successful development of an RPA-LF assay for \u003cem\u003eB. divergens\u003c/em\u003e detection in ticks. The assay demonstrates high sensitivity and specificity under isothermal conditions and provides a portable and reliable tool for field-based molecular surveillance of emerging tick-borne pathogens. Field validation revealed \u003cem\u003eB. divergens\u003c/em\u003e/\u003cem\u003eB. capreoli\u003c/em\u003e in questing ticks from urban green spaces in London, underscoring the importance of this tool for early detection and ongoing surveillance of tick-borne parasites in expanding urban ecosystems.\u003c/p\u003e","manuscriptTitle":"A Recombinase Polymerase Amplification Lateral Flow Assay (RPA-LF) for detection of Babesia divergens","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-03-12 09:46:18","doi":"10.21203/rs.3.rs-8999595/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2026-04-19T21:10:36+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-04-17T19:53:18+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"327493161053474512564765078922831357781","date":"2026-04-07T14:02:13+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-04-07T09:58:08+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"122791854030956505960731386179483027036","date":"2026-03-09T06:07:58+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-03-06T15:19:17+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2026-03-02T12:25:21+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2026-03-02T11:41:16+00:00","index":"","fulltext":""},{"type":"submitted","content":"Parasites \u0026 Vectors","date":"2026-03-01T06:29:39+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"parasites-and-vectors","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"parv","sideBox":"Learn more about [Parasites \u0026 Vectors](http://parasitesandvectors.biomedcentral.com/)","snPcode":"13071","submissionUrl":"https://submission.nature.com/new-submission/13071/3","title":"Parasites \u0026 Vectors","twitterHandle":"@bugbittentweets","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"1831d693-6382-4adf-8c48-574727828c15","owner":[],"postedDate":"March 12th, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"in-revision","subjectAreas":[],"tags":[],"updatedAt":"2026-04-19T21:23:47+00:00","versionOfRecord":[],"versionCreatedAt":"2026-03-12 09:46:18","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8999595","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8999595","identity":"rs-8999595","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}