Dragonfly: A Portable Molecular Diagnostic Platform for Rapid Point-of-Care Detection of Mpox and Other Diseases

doi:10.21203/rs.3.rs-4926828/v1

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Dragonfly: A Portable Molecular Diagnostic Platform for Rapid Point-of-Care Detection of Mpox and Other Diseases

https://doi.org/10.21203/rs.3.rs-4926828/v1

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The explosive global spread of Clade II mpox in humans and sustained human-to-human transmission of the more virulent Clade I mpox in the Democratic Republic of Congo highlight both the current lack of, and critical need for, point-of-care diagnostics to address the spread of this emergent disease. A novel point-of-care molecular diagnostic platform, Dragonfly, was developed that integrates power-free nucleic acid extraction (< 5 minutes) and lyophilized colorimetric LAMP chemistry. The platform achieved an analytical limit-of-detection of 50 genome copies/reaction for monkeypox virus, distinguishing it from other orthopoxviruses, herpes simplex virus and varizella zoster virus. Clinical validation using 164 samples, including 51 mpox positives, demonstrated 96.1% sensitivity and 100% specificity for orthopoxviruses, and 94.1% sensitivity and 100% specificity for monkeypox virus. Our platform provides a rapid, accessible and robust point-of-care solution for monitoring mpox in both low and high resource settings amidst declining worldwide smallpox immunity, with broad potential to the wider field of infectious disease diagnosis as a whole.

Biological sciences/Biological techniques/Lab-on-a-chip

Health sciences/Diseases/Infectious diseases/Viral infection

In 2022, as the SARS-CoV-2 pandemic waned, an unprecedented global mpox outbreak emerged, exhibiting clear epidemiological and clinical differences from historical mpox cases, which were typically associated with zoonotic acquisition and limited human transmission.^1,2 For the first time, sustained human-to-human transmission was observed, primarily through sexual contact, within and between non-endemic and endemic countries.^1,3–6 This outbreak was declared a public health emergency of international concern (PHEIC) by the World Health Organization in July 2022³, rapidly spreading across 117 countries and resulting in 97,208 confirmed cases and 186 deaths.^4,7 Although the origins of the virus remain obscure, an initial and sustained lack of detection is thought to have played a significant role in global dissemination.⁸ The fast-paced global spread of Clade II mpox and its evolving epidemiology⁹ contrasts with the primarily zoonotic Clade I mpox outbreaks that have occurred frequently in Democratic Republic of the Congo (DRC) since the mid 1990s¹⁰. This change underscores the critical importance of early case detection and population surveillance to support effective pandemic preparedness and outbreak management. The importance is further highlighted by the recent description of sustained human transmission of Clade I mpox in eastern DRC, with epidemiology that is strikingly similar to the global Clade II outbreak, and spread by both heterosexual contact as well as men who have sex with men.^10–12 Post hoc investigations into responses to similarly novel and evolving infectious outbreaks, such as HIV^13,14, influenza virus^15,16, Ebola¹⁷, and SARS-CoV-2¹⁸, have consistently placed significant emphasis on the role of diagnostics in successful control, and highlighted the need for solutions suitable for use at the point-of-care (POC) and in low resource settings.¹⁹

The WHO ‘Strategic framework for enhancing prevention and control of mpox 2024–2027’ aims to end human-to-human transmission of mpox to remove the public health and pandemic threat that mpox presents.⁷ The framework requires development of diagnostic assays for use in decentralized sites or POC. The WHO Target Product Profiles for mpox diagnosis^4,7,20 states the importance of differential tests which distinguish multiple diseases, particularly in the prodrome phase when mpox clinical presentation may overlap with other conditions. Distinguishing mpox from similar skin conditions, including chickenpox and herpes simplex, during mpox’s eruptive phase is also critical in identifying cases, informing infection management and reducing onward transmission.^20,21 Preliminary mpox diagnosis is often made clinically, with confirmation by molecular nucleic acid amplification tests (NAATs) or viral culture,^22,23 which severely limits diagnostic solutions in remote and resource limited settings where the endemic disease burden is greatest.^24,25

Protein-based immunoassays such as lateral flow tests (LFTs) offer a rapid, portable, and low-cost solution to POC diagnostics, and are valuable for estimating population-scale infection rates.^26,27 However, numerous retrospective investigations into diagnostic technologies during the COVID-19 pandemic demonstrate that LFTs can have sensitivity limitations, and there is currently no suitably sensitive LFT for mpox.^28–30 Consequently, NAAT methods such as quantitative PCR (qPCR) have remained the gold standard for mpox diagnosis.³¹ Unfortunately, despite excellent specificity and sensitivity, qPCR presents notable limitations, predominantly in POC and near-patient use.³² qPCR often involves complex sample processing, requiring skilled technicians, bulky specialized equipment which is often expensive, and controlled laboratory conditions. All these factors limit portability and restrict deployment in emergencies and in low resource environments, where the current disease burden is greatest and accessible POC diagnostics are most needed.³³

Loop mediated isothermal amplification (LAMP)-based mpox assays can offer an important alternative to qPCR.^34–37 Notably, LAMP is a NAAT approach that operates under isothermal conditions, with the amplification reaction occurring at a constant temperature (typically 60 to 65°C), eliminating the need for a thermocycler and making LAMP a more accessible, cost-effective, and portable option for POC use. Nonetheless, simplifying LAMP-based NAATs for POC use, without compromising accuracy and sensitivity, remains a technological challenge, with sample processing in particular presenting a significant bottleneck.^38–40 Sensitivity has been shown to depend heavily on the quality, purity, and concentration of nucleic acid in the processed sample,^40,41 requiring complex sample preparation protocols to ensure successful amplification. While recent efforts have examined extraction-free, or “direct”, NAATs, to reduce cost and complexity, these have been shown to sacrifice sensitivity through increased amplification inhibition and sample dilution.^42–44

To address these serious diagnostic gaps, we have developed and validated a novel portable molecular sample-to-result POC diagnostic platform called Dragonfly, designed for rapid multi-pathogen detection and differentiation of skin-tropic viruses. Our platform integrates a simple power-free nucleic acid extraction and purification method based on magnetic beads^45,46, combined with a lyophilized colorimetric LAMP technology⁴⁷. This provides a minimally equipment-dependent solution without the need for thermocycling, optical fluorescent readouts, or cold-chain logistics. A sample-to-result kit was developed for the simultaneous detection and differentiation of orthopoxvirus (OPXV) genus, mpox (clades I and II), varicella-zoster virus (VZV), herpes simplex virus type 1 (HSV-1) and, herpes simplex virus type 2 (HSV-2) in a multiplex skin panel, enhancing diagnosis and infection management. The platform’s analytical performance was validated using 164 clinical samples, including 51 mpox clade II positive samples, and was benchmarked against a gold standard extracted qPCR workflow.

1. Platform overview

The platform is comprised of three core components: a single-use sample extraction kit, a lyophilised colorimetric LAMP-based panel, and a low-cost isothermal heat block. (Fig. 1a) In addition, to support the workflow at the POC, the platform includes a sample collection kit, containing a swab and inactivating medium (COPAN eNAT®), a reusable fixed-volume pipette with disposable tips, and an optional tablet with cloud-connected companion software for result logging and centralized data collection. The Dragonfly workflow comprises power-free sample extraction of nucleic acids (< 5 minutes) followed by colour-based molecular detection (< 35 minutes) with visual equipment-free result read-out. A high-level overview is illustrated in Fig. 1b-c, with detailed process images in Fig. 3a-f.

2. Platform components

As described previously,^45,46 the SmartLid technology utilizes a magnetic lid to capture and transfer magnetic nanoparticles and attached DNA/RNA through three simple steps (lysis-binding, wash, and elution) allowing simple, efficient, and power-free nucleic acid extraction without centrifugation or manual pipetting. (Fig. 2) All required buffers are pre-aliquoted in color-coded tubes (utilizing a sequential traffic-light system of red, yellow, and green) and packaged in a cardboard tray along with disposable exact-volume pipettes for sample input and a SmartLid for magnetic nanoparticle manipulation. The fully recyclable carboard packaging also functions as a workstation, facilitating the POC workflow with clear diagrammatic labelling, a space for housing the sample tube, and receptacles for holding the lyophilized test panel during reagent resuspension. (Fig. 3c)

For the test panel, a lyophilized colorimetric LAMP approach was developed to enable long term room temperature storage, visual result readout, and isothermal incubation.⁴⁷ This enables visual identification of positive reactions based on proton production (and subsequent pH drop), which occurs with nucleotide incorporation during DNA polymerase activity. Furthermore, this colour transition, from pink (at high pH) to yellow (at low pH), is compatible with multiple colour vision deficiency (CVD) friendly colour schemes, ensuring maximum usability regardless of user profile.⁴⁸ An off-the-shelf eight-tube PCR strip (4titude®), with individually flip-capped 0.2 mL tubes, was selected to house each reaction, with pre-dispensed and lyophilised master-mix. To enhance traceability, polycarbonate tags, attached in the middle of each tube strip, provide space for data-matrix and alpha-numeric labelling. Combined, these three aspects provide a platform that is as equipment-free as possible, requiring only a low-cost, portable, and user-friendly isothermal heat block that can be powered from mains electricity, battery, or standard 12-volt supply.

3. Cloud-connected companion application

To augment the manual workflow, an optional companion Android application was developed with the goal of improving the user experience and enabling wireless cloud connectivity to integrate with healthcare databases. (Fig. 3g-j) Utilizing the onboard device camera, the application further enhances traceability by scanning data matrices on all consumables at the start of the process. A virtual step-by-step workflow was implemented to guide new users through the process, with progress tracked and displayed, and animated clickable timers provided throughout the workflow to time each step (e.g., buffer shaking). Test panel incubation is also monitored and timed, with assistance provided to remind the user of the incubation status of each sample and their location in the heat block. After incubation, result interpretation and recording were augmented through an image capture process, in which a cropped and enlarged view of the eight tubes is presented with buttons to select observed colours. Depending on the specific panel scanned in at the beginning of the process, results are then interpreted and displayed. All results, along with captured test panel images, are automatically stored and summarized in an AWS cloud-accessible dashboard for export, review, and traceability.

4. Skin Panel Design

Our test panel was designed to target orthopoxviruses (OPXV), monkeypox virus (MPXV), varicella-zoster virus (VZV), herpes simplex virus type 1 (HSV-1), and herpes simplex virus type 2 (HSV-2). VZV, HSV-1 and HSV-2 can cause skin rashes and lesions confounding MPXV diagnosis. In addition, zoonotic OPXV like cowpox or borealpox virus have been associated with fatal human infections in Europe and America.⁴⁹ Therefore, our panel aims to facilitate the correct diagnosis of mpox cases and capture the emergence of zoonotic OPXV species. The layout of the panel is shown in Fig. 1c and includes the following targets from left to right: colour control (CC), OPXV, MPOX, VZV, HSV-1, HSV-2, extraction control (EC), and internal control (IC). Two LAMP assays were included per lyophilized reaction mix for OPXV, MPOX and VZV, and one assay each for HSV-1 and HSV-2. In the case of the orthopoxvirus genus, genes E9L (viral DNA polymerase gene, conserved gene segment within all Eurasian OPXVs) and F13L (encoding a conserved protein in OPXVs required for viral maturation and release from the infected cell⁵⁰) were selected, with the first one also used as target by the US Centers for Disease Control (CDC) qPCR assay⁵¹. For MPXV, specific assays targeting conserved intraspecies regions of genes G2R (encoding a viral tumour necrosis factor receptor) and A9L (VACV orthologue that encodes a morphogenesis factor) were designed, in order to cover both clade I and II without cross-reactivity with other OPXV. G2R is currently used as the target of choice by the CDC qPCR assay.⁵² For the third viral target, an assay was designed targeting ORF28 gene of VZV, and used in conjunction with another assay from Okamoto et al.⁵³ targeting ORF62 gene. For the specific detection of HSV-1, an assay from Kaneki et al.⁵⁴ targeting the UL1 gene was utilized and for HSV-2 the assay was designed targeting the US4 gene.

The panel includes three additional control reactions to ensure reliable results. First, as Dragonfly utilizes a pH-based colorimetric indicator, variations in sample and extracted elution pH were observed to affect the starting resuspended (negative) reaction colour. To account for this, a “colour reference control” was added (which excludes amplification enzymes), ensuring that one tube would always remain pink, and thus represent a pH-adjusted negative colour. Next, an “internal control” reaction mix was added, which contains the DNA template specific to its assay, ensuring amplification under ideal operating conditions. This control acts as a confirmatory reaction to indicate that the test panel is in good working order (e.g., not damaged due to improper storage, incubated at the wrong temperature, or excessively inhibited). Finally, a “human extraction control” was included, which targets the human housekeeping beta-actin gene⁵⁵], providing a confirmation that the extraction process performed correctly and that the sample source was sufficiently swabbed. A valid test result, whether positive or negative, requires that the three described control reactions are pink (negative), yellow (positive), and yellow (positive) respectively.

5. Workflow

The extraction and detection workflows were optimised to achieve a balance between protocol complexity, protocol time, and performance: Inactivated swab eluent (400 µL) was transferred from the sample collection tube into Tube A (containing lysis/binding buffer and magnetic beads) using a disposable exact volume pipette. (Fig. 3a) The included SmartLid was then used to transfer magnetic beads and attached nucleic acids through a series of three sample extraction steps (A: lysis, B: wash, and C: elution). (Fig. 3b) At each step, resuspended magnetic beads were mixed with the buffers for 30 seconds through manual shaking without the magnet, followed by insertion of the magnet to collect the magnetic beads, and a 30 second drying step to evaporate any remaining solvents prior to elution. Once nucleic acids were eluted into Tube C, the magnetic beads were removed with the SmartLid. Next, a 20 µL fixed volume pipette (using a disposable tip) was used to transfer elution into each tube of the test panel, resuspending the lyophilized reagents. (Fig. 3c) Unlike more expensive laboratory micro-pipettes, the included fixed-volume pipettes were modified to remove the secondary ‘blow-out’ stage, simplifying the pipetting process for novice users without significantly reducing accuracy. Once all caps are firmly closed, test panels were placed into one of the four heat block rows (Fig. 3d) and incubated for 35 minutes at 63.5˚ C (heater locked at temperature to reduce potential user error). After incubation, test panels were placed into a result capture card, where developed colours could be compared to a key directly below each tube, indicating the result: pink for negative, yellow for positive. (Fig. 3e-f) The entire process, from sample-to-result, was optimised to take < 40 minutes per sample, with each subsequent sample able to be extracted while previous samples were being incubated, yielding a continuous throughput of > 12 samples per hour per user.

6. Analytical sensitivity and specificity of LAMP assays

Sequences of selected target genes were collated from NCBI, and primer sequences are shown in Table S1. Analytical sensitivity was evaluated using serial dilutions of synthetic DNA (one for each target), and the obtained standard curves had a correlation (R²) of 80 to 99% (shown in Figure S1). Time-to-positive (TTP) values across all tested concentration ranging from 10¹ to 10⁷ copies per reaction were within 15 min except for HSV-2 that was 25 min. All assays had an overall limit of detection of 10 to 500 copies per reaction as shown in Fig. 4b and Table S2. Analytical specificity was assessed in-silico based on sequence alignments and experimentally using extracted nucleic acids from commercially available viral particles (Vircell, MBTC032-R, Zeptometrix CATALOG# NATHSV-6L and Zeptometrix CATALOG# NATVZV-STQ). As shown in Fig. 4c, the LAMP assays only amplified their specific target.

7. Sample-to-result evaluation with viral particles

Commercially available MPXV and HSV-1/HSV-2/VZV viral particles (Vircell, MBTC032-R and MBTC016) were spiked into eNAT® inactivation buffer at various concentrations as shown in Table S3. Extractions were performed using Dragonfly Sample Preparation Kits, with eluted nucleic acids then used to resuspend our Skin Infection Viral Test Panels which were then incubated for 35 minutes. The limit-of-detection (LOD) for OPXV and VZV was determined to be 1.25×10³ copies per mL which translates to 50 copies per reaction assuming 100% extraction efficiency. The LODs for MPXV and HSV-1 were determined to be 2.5×10³ copies per mL, and 3.5×10³ copies per mL for HSV-2. Assuming 100% extraction efficiency, this corresponds to 100 and 143 copies per reaction for MPXV/HSV-1 and HSV-2, respectively. Examples of our Skin Infection Viral Test Panels results are shown in Fig. 4e. Lastly, specificity of the panel was further confirmed using cowpox (CPXV) and vaccinia (VACV) viral particles at concentrations of 5×10⁵ PFU per mL, where appropriately only the OPXV reaction turned yellow, validating the specificity of the panel (Fig. 4e).

8. Clinical validation from skin swabs

164 surplus extracts from skin swabs, which had been submitted to North West London Pathology (NWLP) for testing as part of clinical care, were utilized to determine if the described viral DNA targets could be detected in clinical samples with the Dragonfly platform. Results were compared to identifications obtained using gold standard automated nucleic acid extraction and real-time multiplex qPCR detection. The samples, collected in Roche COBAS PCR media (P/N: 07958030190) included 51 mpox Clade II positive samples and 40 samples positive for one or more herpes viruses. Our platform demonstrated high analytical performance, with 96.1% (95% CI of 86.5–99.5%) sensitivity and 100% (95%CI of 96.8–100%) specificity for OPXV, and 94.1% (95%CI of 83.8–98.8%) sensitivity and 100% (95%CI of 96.8–100%) specificity for mpox. Distributions of the C_t values of positive samples are shown in Fig. 4d. There were two false negatives for OPXV and mpox, both of which had qPCR C_t values above 33 C_t (34.37/33.61 C_t and 35.96/34.97 C_t, for OPXV and mpox respectively) and one false negative exclusively for mpox with a qPCR Ct value of 28.65 C_t.

Swabs included 10 VZV, 20 HSV-1 and 10 HSV-2 infections that had been identified previously by qPCR, representing 24.4% (40/164) of the samples analysed and including 4/164 (6.3%) co-infections with mpox (three with HSV-1, and one with HSV-2). Dragonfly detected 9/10 VZV, 18/20 HSV-1, and 7/10 HSV-2 cases (85% of the tested positive samples), including three out of the four co-infections. Further details are shown in Figure S2. All the samples resulted in a positive human extraction control reaction (targeting the beta-actin gene) using our platform, verifying the quality of the extraction and that the sample was correctly collected, limiting potential for false negatives due to user error or insufficient sampling. To verify results of the human extraction control reaction, a TaqMan assay targeting gene RnaseP was used for the qPCR comparator, with a distribution of C_t values shown in Figure S3.

Mpox, an infection caused by MPXV, is historically associated with localised outbreaks in endemic countries of central, west and east Africa, with limited human-to-human transmission. In 2022 a global mpox Clade II outbreak was declared a public health emergency of international concern (PHEIC) by the WHO. The rapid global spread of and altered epidemiology of this outbreak⁹ highlighted the importance of early case detection and population surveillance to support effective pandemic preparedness and management.

In this study, we developed and validated a novel POC molecular diagnostic system, Dragonfly, for the accurate, simultaneous differential detection of mpox and other associated skin tropic virus diseases to support the differential diagnosis of mpox from OPXV, HSV-1, HSV-2 and VZV, all of which present with similar skin lesions. Our platform combines simple power-free nucleic acid purification^45,46 with patented lyophilized colorimetric LAMP technology⁴⁷ to achieve PCR grade analytical performance away from centralised laboratories, without, specialist laboratory equipment, cold chain or skilled laboratory personnel. Dragonfly significantly reduces the reliance on laboratory-based equipment, forgoing complex optical fluorescent systems and thermocyclers, and enabling use in remote and resource limited settings. Our Skin Infection Viral Test Panel, which incorporates MPXV, OPXV, VZV, HSV-1, and HSV-2 targets was validated using 164 clinical skin lesion samples, including 51 mpox Clade II positive samples, 10 VZV, 20 HSV-1 and 10 HSV-2 positive samples. The results were compared to gold standard automated extraction and TaqMan-based qPCR assays, showing high sensitivity and specificity.

The study was limited, however, by the low number of available confirmed HSV-1, HSV-2 and VZV samples. This was a result of the samples being collected at the height of the mpox outbreak and thus during a temporary decline in HSV/VZV testing. A wider validation for these targets should be examined as part of future studies. Additionally, our system was designed to extract samples from eNAT® inactivation buffer, while all collected samples for this study were stored in Roche COBAS PCR media. Accordingly samples were diluted 1:2 in eNAT® inactivation buffer to remain consistent with the intended use of the platform. Should samples be collected from patients and directly stored in eNAT® in future studies, the LOD would be expected to improve proportionally to the dilution factor above. Furthermore, all clinical mpox positive samples examined were Clade II, and collected from the same outbreak, further work will be required to understand the analytical performance on a broader strain collection. While preliminary in-silico analysis and experimental testing with synthetic DNA (Figure S1) confirmed that our assays cover Clade I, further work will nonetheless be required to confirm performance in a clinical setting. Finally, although the panel was designed for this study to include mpox Clade I and II in the same reaction (i.e. to not differentiate clades I and II), it would be a simple matter to reconfigure the panel (for example, by instead combining the HSV-1 and HSV-2 assays into one reaction or include additional targets such as molluscum contagiosum which can cause lesions similar to mpox) to differentiate mpox clades I and II, should that be deemed necessary.

The infectious disease landscape is dynamic, marked by constant changes in pathogen behaviour, emerging threats, and shifts in environmental and socioeconomic factors requiring diagnostic platforms to adapt.⁵⁶ Recent data describes a growing Clade I mpox outbreak that is spreading in a new epidemiologic pattern analogous to that of the global Clade II outbreak. Contrary to enclosed and complex microfluidic cartridges used in many other POC diagnostic systems,⁵⁷ Dragonfly can responsively adapt to emerging needs. It has an easy to manufacture design, with extraction and amplification reagents that are rapidly swappable and customizable with generic off-the-shelf packaging solutions. This create an adaptable system that can be repurposed for different sample types (e.g., blood, bacterial, stool, etc.), targets (e.g., viral, bacterial, etc.), and applications in response to emerging requirements. For example, the Skin Infection Viral Test Panel described in this study was rapidly adapted into a dual-sample two-pathogen panel for higher throughput processing and reduced costs (Fig. 5). Alongside an updated companion application suitable for personal mobile phone use (Fig. 5a), a dual-sample vortex tool was developed (Fig. 5b-d) to augment the already rapid diagnostic workflow. (Associated training video can be found at www.youtube.com/watch?v=Oz1kvwJyNzQ.)

The robustness of a diagnostic test is a key enabler for its accessibility and utility in resource-limited settings.^58,59 Traditional molecular tests often require stringent transport and storage conditions, such as refrigeration, to maintain stability of their many temperature sensitive reagent components, creating logistical challenges.⁶⁰ In contrast, the our platform utilizes novel lyophilized colorimetric LAMP technology⁴⁷, which enables room temperature shipping and storage, ensuring the adaptability of the diagnostic platform to diverse environmental conditions, including where cold chain logistics are not available. Moreover, the compact and lightweight nature of the Dragonfly platform, weighing less than 1.5 kilograms (including the isothermal heat block) and fitting in a backpack, facilitates on-the-go testing and allows healthcare professionals to serve remote or underserved areas efficiently.

In the years since the publication of the original ASSURED (affordable, sensitive, specific, user-friendly, rapid, equipment-free, deliverable) criteria for POC diagnostics in resource limited settings connectivity and technological integration have become more widespread and provide opportunities for improved real-time diagnosis, surveillance and monitoring.⁶¹ A user-friendly companion app was therefore developed with cloud-based data storage and a dashboard for result visualisation, real-time data integration and user/result quality assurance data. Such connectivity solutions not only increase quality assurance for POC tests, but also allow for centralized and real-time decision-making, even across tiered laboratory systems and during outbreak investigations and global health emergencies.

Finally, cost-effective diagnostic platforms can be game-changers in the landscape of molecular sample-to-result systems. The primary equipment required by the majority of portable molecular platforms can cost tens of thousands of pounds. This high price point is a significant barrier for accessibility and widespread adoption, limiting the reach of molecular platforms across healthcare sectors and geographical locations. In contrast, the simple isothermal heater, the most expensive component of our system, is manufacturable for <£100, enhancing the accessibility and economically viability on a large scale.

As emphasised by the recent WHO Strategic framework for enhancing prevention and control of mpox (2024–2027)⁷, there is a need for POC solutions that can support early mpox detection and disease surveillance, especially in resource limited settings where access to diagnostics is more restricted.^62,63 Although there has been an increase in the adoption of isothermal chemistries such as LAMP^64–66 or RPA in combination with CRISPR⁶⁷, they are still primarily laboratory-based, requiring trained personnel, sample transport to centralised facilities, expensive equipment and a cold chain. The Dragonfly platform’s accuracy, portability, and rapid multi-pathogen sample-to-result capability make it a versatile tool with significant potential to contribute to global efforts in combating both emerging and endemic infectious diseases, particularly in low-resource environments.

9. LAMP assay design

LAMP assays were designed using the software Primer Explorer version 5.0 (url{http://primerexplorer.jp/lampv5e/index.html), based on sequences retrieved from NCBI GenBank database (url{https://www.ncbi.nlm.nih.gov/genbank/). Sequences were aligned using the MUSCLE algorithm⁶⁹ in Geneious 10.0.5 software and primers were designed to specifically detect the target DNA without cross-reactivity. All primers were purchased from Integrated DNA Technologies (IDT) and rehydrated in nuclease-free buffer at 400 µM. The LAMP assay specific to the human house-keeping gene beta-actin⁵⁵, LAMP-ACTB, was previously described and the LAMP assay specific to HSV-1 and HSV-2, were also previously described^70,71. All the primer sequences used in this study are shown in Table S1.

10. LAMP reaction conditions

Fluorescent detection: LAMP reactions were carried out at a final volume of 10 µL per reaction. Each mix contained the following: 1 µL of 10× custom isothermal buffer (pH 8.5–9), 0.5 µL of MgSO4 (100 mM stock), 0.56 µL of dNTPs (25 mM stock), 0.3 µL of BSA (40 mg/mL stock), 1 µL of 10× LAMP primer mix (F3/B3 2.5 µM, LF/LB 10 µM and FIP/BIP 20–40 µM), 0.25 µL of Syto9 dye (20 µM stock), 0.1 µL of NaOH (0.5 M stock), 0.04 µL of Bst 2.0 WarmStart DNA polymerase (120K U/mL stock), 1 µL sample, and enough nuclease-free water (Invitrogen) to bring the volume to 10 µL. Reactions were loaded into 96-well plates and were performed at 63 ⁰C during 35 min using a QIAquant real-time PCR (QIAGEN) and LightCycler 96 (LC96) instrument (Roche). One melting cycle was performed at 0.1 ⁰C/s from 63 ⁰C up to 97 ⁰C for validation of the specificity of the amplified products. Non-template control (NTC) was included in every experiment.

Colorimetric detection: LAMP reactions were performed using the colorimetric Dragonfly Skin Infection Viral Test Panels (ProtonDx) at a final volume of 20 µL adding 20 µL of sample elution in nuclease-free water. Amplification reactions were carried out in a portable thermal block (Proton Dx) during for 35 min at 63.5 ⁰C.

11. Analytical sensitivity and specificity of LAMP assays

Analytical sensitivity was evaluated using 10-fold serial dilutions of synthetic DNA (gBlocks from Integrated DNA Technologies) ranging from 10⁷ to 10 copies per reaction with half-dilutions between 10³ and 10 copies per reaction (i.e., 500 and 50 copies per reaction). Dilutions were performed using nuclease-free water. Each condition was run in triplicate using a LC96 instrument.

Analytical specificity was performed in-silico based on sequence alignments and mapping of primers, and experimentally using extracted DNA from viral particles including AMPLIRUN® TOTAL MONKEYPOX VIRUS CONTROL (SWAB) from Vircell (reference MBTC032-R), HSV1&2 Positive control from ZeptoMetrix (CATALOG# NATHSV-6L) and Varicella-Zoster Virus Stock (Quantitative) from ZeptoMetrix (CATALOG# NATVZV-STQ). Extractions were performed manually using QIAamp Viral RNA Mini Kit as recommended by the manufacturer using 140 µL as input.

12. Statistical analysis and ethics

Use of fully anonymized surplus samples was approved by the West London National Research Ethics Committee (approval no. 06/Q0406/20). Sample size has been calculated based on the equation reported by Banoo et al.⁷² Considering a marginal error of 10% with a CI of 95%, and an estimated 90% sensitivity and specificity, the size of the study population will be 35 and we included 164 samples. Time-to-positive (TTP) data is presented as mean TTP ± standard deviation.

13. Extraction of nucleic acids from skin swabs and PCR reaction conditions

Nucleic acids were extracted from clinical samples as part of routine diagnostic service at North West London Pathology using Qiagen Viral RNA kit for the Qiagen EZ-1 advanced Excel system, and were tested with an MPOX laboratory developed test (LDT) as described in Pond et al.⁷³ Briefly, this consisted of a TaqMan assay targeting G2R WA for MPOX clade II⁷⁴, and a TaqMan assay targeting E9L gene of OPXV⁷⁵. The MPOX LDT was reported as “MPOX detected” if amplification occurred in both the G2R WA and E9L-NVAR assays. Samples were reported as MPOX Indeterminate if amplification was observed to occur in only one of the two MPOX LDT qPCR assays.

For this study, lesion swabs collected in Roche COBAS PCR media (P/N: 07958030190) were archived at -20ºC prior to testing. Roche COBAS PCR media was diluted in eNAT® prior to extraction, using a 1:2 ratio (sample in COBAS PCR media:eNAT®). The 164 samples were extracted using the QIAsymphony® DSP Virus/Pathogen Midi Kit (Cat. No. 937055) in combination with the QIAsymphony SP instrument. Briefly, the protocol used was Complex 400 which process 400 µL of sample as input. Elutions were performed in cooled 96 low-skirt plates with a final volume of 110 µL per sample. Elutions were tested on the day and then stored at -80 ⁰C. Published TaqMan assays⁷⁶ as detailed in Table S3, were used to detect the presence of VZV, HSV-1, and HSV-2. The Promega GoTaq Probe qPCR (Cat. No. A6102) was used at a final volume of 20 µL per reaction following this protocol: 2x GoTaq qPCR mastermix (10 µL), 10x assay mix (2 µL), 5 µL of sample, and enough nuclease-free water. The assay mix final concentration was forward (400 µM), reverse (400 µM) and probe (200 µM). The cycling conditions were: 1 cycle at 95 ⁰C for 2 min, and 45 cycles at 95 ⁰C for 3 s followed by 60 ⁰C for 30 s. Experiments were performed using a QuantStudio5 instrument reading in FAM channel. Published human control assay targeting RNaseP from the CDC was used to verify the human origin of the sample and the quality of the extraction. The Promega GoTaq Probe 1-step RT-qPCR (Cat. No. A6121) was used at a final volume of 20 µL per reaction following this protocol: 2x GoTaq qPCR mastermix (10 µL), GoScript RT Mix (0.4 µL), assay ready mix (1.5 µL), 5 µL of sample, and enough nuclease-free water. The cycling conditions were: 1 cycle at 45 ⁰C for 15 min, 1 cycle at 95 ⁰C for 2 min, and 45 cycles at 95 ⁰C for 3 s followed by 55 ⁰C for 30 s. Experiments were performed using a QIAquant instrument reading in FAM channel.

14. Inactivation of skin swabs using Dragonfly sample preparation

eNAT® buffer (COPAN) was tested neat against VACV WR or diluted 1/10 with PBS against HSV-1. 1 ml eNAT® buffer was aliquoted into screwcap tubes for the virus (VC) and cytotoxicity (CC) controls, and triplicate screwcap tubes for exposure to virus. 10 µl cell culture media was added to the cytotoxicity control. 10 µl virus was added to the triplicate tubes as well as the virus control tube, and incubated at room temperature for the indicated length of time. After the contact time, each sample was serially diluted in cold DMEM supplemented with 2% v/v FCS and 1% v/v pen/strep. Additionally, a neutralisation control (NC) was set up to confirm that virucidal activity was halted after the contact time by dilution. First, the cytotoxicity control was diluted 1/9 in cold DMEM and allowed to neutralise by mixing. The virus control was then diluted 1/10 in the same tube, before continuing with the serial dilution.

Confluent monolayers of BSC40 (for VACV titration) or VERO (for HSV-1 titration) cells were used to perform the plaque assay in 6-well plates. Cell culture media was removed from the cell monolayers and 0.5 ml diluted samples was plated in duplicate, and allowed to adsorb for 1 hour at 37°C, with rocking after 30 minutes. The inoculum was then removed and a semi-solid overlay was added to each well (1.5% CMC, 1x MEM), before incubation at 37°C for 2–3 days, or until plaques were countable.

After incubation, the overlay was removed and the wells were washed carefully with PBS. Enough crystal violet (1% crystal violet, 70% ethanol) was added to cover the base of each well and incubated at room temperature for at least 30 minutes before excess stain was removed and the wells rinsed with water. Plaques were counted and the titer for each sample was calculated.

15. Dragonfly system Workflow

The Dragonfly workflow comprises, (i) nucleic acid extraction with the Dragonfly Sample Preparation Kits and (ii) pathogen detection with the Dragonfly Skin Infection Viral Test Panel. First, nucleic acid extraction buffer compositions, volumes, mixing durations, and total number of steps was substantially based on original SmartLid method for viral nucleic acid extraction.^45,46 However, due to the cited method being developed for the extraction of SARS-CoV-2 RNA from inactivating storage media (COPAN eNAT®), the protocol was further optimized to improve nucleic acid (DNA) yield from clinical isolates that were stored in non-inactivating media (Roche COBAS PCR Medium). Samples were diluted at a 1:2 ratio (sample in ROCHE COBAS PCR Medium:eNAT®) and 400 µL were used as input for the nucleic acid extraction. The guanidinium thiocyanate (GTC) and magnetic bead (TurboBeads) concentrations were increased for the lysis buffer (Tube A), and two wash steps were utilized instead of one. Next, volumes of 20 µL were utilized for each lyophilized colorimetric LAMP reaction to strike a balance between reagent cost, sensitivity, and ease of user liquid handling. Finally, a sufficient reaction duration of 35 minutes was determined through observing reaction colors at 5-minute intervals during the LOD experiments. This incubation duration was also confirmed to not yield non-specific amplification through negative control experiments that were incubated for up to 50 minutes at 63.5°C.

Funding

This work was supported by the Department of Health and Social Care-funded Centre for Antimicrobial Optimisation (CAMO) at Imperial College London; the Imperial College London UKRI Impact Acceleration Account [MRC – MR/X502959/1]; the Wellcome Trust CAMO-Net programme [226691/Z/22/Z]; Wellcome Trust Innovator Award [215688/Z/19/Z]; the Biotechnology and Biological Science Research Council [BB/X011569/1]; the Engineering and Physical Sciences Research Council [grant ref: EP/T51780X/1] and the Imperial College Research Fellowship [WDPI.G09074]. In addition, this research was funded by the NIHR [NIHR134694] using UK international development funding from the UK Government to support global health research. The views expressed in this publication are those of the authors and not necessarily those of the NIHR or the UK government. The authors acknowledge the support of the local diagnostic laboratory (Northwest London Pathology) and the NIHR Biomedical Research Centre awarded to Imperial College London. AH., SS, PG and JRM are affiliated with the NIHR Health Protection Research Unit (HPRU) in Healthcare Associated Infections and Antimicrobial Resistance at Imperial College London in partnership with the UK Health Security Agency, in collaboration with, Imperial Healthcare Partners, the University of Cambridge and the University of Warwick. The views expressed in this publication are those of the authors and not necessarily those of the NHS, the National Institute for Health Research, the Department of Health and Social Care, or the UK Health Security Agency. AH is a National Institute for Health Research (NIHR) Senior Investigator.

Author Contributions

MLC and KMC authors contributed equally to the work. Study concept and design: MLC, KMC, MP, and JRM. Acquisition, analysis, or interpretation of data: All authors. Drafting the manuscript: MLC, KMC, DU, CMM and JRM. Critical revision of the manuscript: All authors. The manuscript was written through the contributions of all authors. All authors have given approval to the final version of the manuscript.

The authors declare the following competing financial interest(s): MLC, KMC, IP, SM, NM, MC, KMS, OWS, PG and JRM have financial interest on ProtonDx Ltd, which currently has exclusive license to intellectual property linked to Dragonfly and its associated trademark. All authors declare that they have no other conflict of interest related to this work. All the authors declare that they do not have any other known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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Yes there is potential Competing Interest. MLC, KMC, IP, SM, NM, MC, KMS, OWS, PG and JRM have financial interest on ProtonDx Ltd, which currently has exclusive license to intellectual property linked to Dragonfly and its associated trademark. All authors declare that they have no other conflict of interest related to this work. All the authors declare that they do not have any other known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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Dragonfly: A Portable Molecular Diagnostic Platform for Rapid Point-of-Care Detection of Mpox and Other Diseases

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Abstract

Figures

Introduction

Results

1. Platform overview

2. Platform components

3. Cloud-connected companion application

4. Skin Panel Design

5. Workflow

6. Analytical sensitivity and specificity of LAMP assays

7. Sample-to-result evaluation with viral particles

8. Clinical validation from skin swabs

Discussion

Online Methods

9. LAMP assay design

10. LAMP reaction conditions

11. Analytical sensitivity and specificity of LAMP assays

12. Statistical analysis and ethics

13. Extraction of nucleic acids from skin swabs and PCR reaction conditions

14. Inactivation of skin swabs using Dragonfly sample preparation

15. Dragonfly system Workflow

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References

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