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. Author manuscript; available in PMC: 2019 Nov 30.
Published in final edited form as: Biosens Bioelectron. 2018 Aug 10;120:93–101. doi: 10.1016/j.bios.2018.08.005

Integrated LAMP and immunoassay platform for diarrheal disease detection

C R Phaneuf 1, B Mangadu 2, H M Tran 3, Y K Light 4, A Sinha 5, F W Charbonier 6, T P Eckles 7, A K Singh 8, C-Y Koh 9
PMCID: PMC6145809  NIHMSID: NIHMS1505658  PMID: 30172236

Abstract

The challenges of diagnosing infectious disease, especially in the developing world, and the shortcomings of available instrumentation have exposed the need for portable, easy-to-use diagnostic tools capable of detecting the wide range of causative microbes while operating in low resource settings. We present a centrifugal microfluidic platform that combines ultrasensitive immunoassay and isothermal amplification-based screening for the orthogonal detection of both protein and nucleic acid targets at the point-of-care. A disposable disc with automatic aliquoting inlets is paired with a non-contact heating system and precise rotary control system to yield an easy-to-use, field-deployable platform with versatile screening capabilities. The detection of three enterotoxins (cholera toxin, Staphylococcal enterotoxin B, and Shiga-like toxin 1) and three enteric bacteria (C. jejuni, E. coli, and S. typhimurium) were performed independently and shown to be highly sensitive (limit of detection = 1.35–5.50 ng/mL for immunoassays and 1–30 cells for isothermal amplification), highly exclusive in the presence of non-specific targets, and capable of handling a complex sample matrix like stool. The full panel of toxins and bacteria were reliably detected simultaneously on a single disc at clinically relevant sample concentrations in less than an hour. The ability of our technology to detect multiple analyte types in parallel at the point-of-care can serve a variety of needs, from routine patient care to outbreak triage, in a variety of settings to reduce disease impact and expedite effective treatment.

Keywords: microfluidics, centrifugal, point-of-care, immunoassay, isothermal amplification, LAMP, enteric pathogens

1. Introduction

Diagnostic methods must evolve to meet the need for accurate, timely, and comprehensive screening of infectious diseases. In addition, delivering these capabilities at the point-of-care is critical in settings that lack access to laboratory infrastructure. Despite the many innovations in molecular biology and instrumentation, the versatility of state-of-the-art devices remains limited. Since symptoms can often be traced to a variety of potential etiologies, including viruses, parasites, bacteria, and the toxins they produce, a comprehensive diagnostic tool should be able to detect targets in more than one of these categories. While methods of multiplexing have been developed for more efficient panel-based testing of a single class of target, the flexibility to probe multiple target classes has eluded the single-purpose devices of the past and present.

Traditionally, three techniques have been the gold standard methods of identifying pathogens: culture and microscopy-based techniques (Zhao et al., 2014) , nucleic acid amplification tests (NAATs), and immunoassays. Time from sample to answer from these processes can vary from several hours, in the case of NAATs and immunoassays, to several days , in the case of culture and microscopy-based detection, and all require a well-equipped laboratory staffed with experienced technicians. Although follow-up patient evaluations often still require conventional culture methods for test-of-cure purposes, NAATs and immunoassays occupy an important role in medical diagnostics.

Chief among NAATs is the polymerase chain reaction (PCR), which can provide highly sensitive and specific detection in hours . Variants of this technique include quantitative PCR, which provides real-time detection and quantification of a genetic target, and multiplex PCR, which simultaneously amplifies multiple distinct targets for efficient screening of a sample. Another type of NAAT is isothermal amplificationwhich is performed at a single temperature, allowing for simpler, more energy-efficient instrumentation. Of the isothermal methods available, loop-mediated isothermal amplification (LAMP) has established itself as a robust diagnostic tool, providing performance comparable to PCR methods while tolerating relatively high amounts of contaminants and therefore requiring less sample preparation (Mori and Notomi, 2009; Song et al., 2005; Yamazaki et al., 2009).

While most of the recent advances in enteric pathogen screening have been nucleic acid-based, immunoassays can be extremely sensitive and allow for the direct identification of various toxins. This can be critical to differentiate between toxigenic and non-toxigenic strains of bacteria in endemic areas where incidence is low or in the early stages of an outbreak (NCID and PAHO, 1994). Methods such as enzyme-linked immunosorbent assay (ELISA) are commonplace in sophisticated laboratories, while the more recently developed dipstick tests, also referred to as immunochromatographic, lateral flow, or rapid diagnostic tests, can operate instrument- offering simplicity and ease-of-use (Fleece et al., 2016; Khan et al., 2016; Muldoon et al., 2007; Ontweka et al., 2016; Page et al., 2012; Zhao et al., 2016). Despite these promising features, the sensitivity of dipstick assays cannot currently compete with most NAATs and therefore fail to detect low but clinically relevant analyte concentrations.

Most of these technologies are limited to detecting a single class of target molecule. One exception is the Luminex LabMAP system, which uses PCR and nucleic acid hybridization and a capture-sandwich immunoassay to detect both DNA and protein targets, respectively, with labeled microspheres (Dunbar et al., 2003). Despite successfully demonstrating the possibility of parallel detection, the method suffers from relatively insensitive limits of detection and requires significant time and labor investment for sample preparation. Another microparticle-based approach known as the bio-barcode assay obviates the need for PCR and reports high sensitivity detection of DNA and protein targets but still requires substantial reagent preparation (Hill and Mirkin, 2006).

These modern detection strategies have vastly improved the speed, accuracy, and sensitivity of medical diagnostics, but their impact has been primarily limited to centralized laboratories with the budgets and infrastructure to operate the myriad benchtop instruments used to conduct these tests. To address the problem of widespread infectious disease in the regions most affected, detection tools must be field-deployable and capable of operating in resource-limited settings. A useful rubric for such diagnostics is the ASSURED criteria: affordable, sensitive, specific, user-friendly, rapid and robust, equipment-free and deliverable to end-users (Peeling et al., 2006; St John and Price, 2014). Hence the promise of microfluidics, which aim to reduce the cost, complexity, and turnaround time of analysis by miniaturizing and integrating established diagnostic techniques as well as forging novel methodologies (Sackmann et al., 2014; Yager et al., 2006). A subset of the field known as lab-on-a-disc or centrifugal microfluidics offers many unique advantages for the development of simple, portable devices (Smith et al., 2016; Tang et al., 2016).

The need for more portable and versatile diagnostic tools is clear for the case of diarrheal disease. Reports from the World Health Organization (WHO) indicate that diarrheal disease is the second leading cause of death in children under the age of five. Of the approximately 2 billion cases occurring annually, diarrhea kills over one million children (Wang et al., 2016). Although diarrhea represents a major cause of morbidity and a significant health care burden in developed regions, the most devastating impact occurs in the developing world, where a combination of socioeconomic and cultural factors conspire to produce a shockingly high concentration of these preventable deaths (Farthing et al., 2013). There is great potential to curtail the burden of diarrheal disease through increased accessibility and adoption of effective detection technologies capable of operating in low-infrastructure settings (Caliendo et al., 2013).

The work presented here offers an alternative to existing methods with unique advantages of portability, ease-of-use, and versatility. Specifically, our platform harnesses the benefits of miniaturization for portability and introduces a new method of combining two modes of detection into a single step for more comprehensive screening. This technology has its foundations in previous efforts toward demonstrating high-sensitivity biological detection using a novel sedimentation-based immunoassay on a centrifugal microfluidic platform (Koh et al., 2015; Phaneuf et al., 2016; Schaff and Sommer, 2011). With the addition of a non-contact temperature control system and an easy-to-use microfluidic disc design, we have developed a field-deployable platform capable of simultaneously detecting both nucleic acid and protein targets. We demonstrate this orthogonal detection strategy by performing an array of immunoassays and LAMP reactions on a single disc for an enteric panel consisting of three toxins: cholera toxin, Staphylococcal enterotoxin B (SEB), Shiga-like toxin 1 (Stx1) and three bacteria: Campylobacter jejuni, enterohemorrhagic Escherichia coli (EHEC, specifically O157:H7), and Salmonella typhimurium, which will be abbreviated henceforth as C. jej, E. coli, and S. typ. These targets represent the causes of some of the most common foodborne and waterborne illnesses. In addition to performing co-detection of toxins and bacteria, the sensitivity, specificity, and sample matrix independence of the assays are evaluated to comprehensively demonstrate the efficacy of the system.

2. Methods and materials

2.1. Instrument

The centrifugal microfluidic platform developed for this study is composed of three subsystems: rotary control system, optical detection system, and heating system. These subsystems work in concert to provide the functionalities of loading and centrifugation, fluorescence-based detection, and temperature control for isothermal amplification chemistries.

The rotary control system consists of a brushless DC motor with a 12-bit magnetic absolute encoder (Faulhaber) paired with a reflective object sensor (Optek), also known as an optical switch. The brushless motor allows for both precise velocity control of the microfluidic disc, operating at a wide range of speeds from approximately 5 to 5000 RPM for sample loading and centrifugation, and position control for moving incrementally between samples for data collection. An aluminum hub coupled to the motor shaft locates the microfluidic disc and a thumbscrew secures the disc to the hub. The optical switch interacts with an etched marking on the disc to detect the home position, which corresponds to the location of the first sample chamber. To carry out endpoint fluorescence detection of a disc, home is first located, then the encoder tracks the position of the disc as it incrementally moves across each chamber, sweeping each of the chamber tips at a speed of 5 RPM while measuring fluorescence at 1 kHz.

The optical detection system is configured for high sensitivity laser-induced fluorescence (LIF) measurement. The key elements include a 635 nm red laser diode (Edmund Optics), photomultiplier tube (PMT) module (Hamamatsu), and a filter set (Semrock) consisting of a 640/14 nm excitation filter, 676/29 nm emission filter, and dichroic beamsplitter with an edge wavelength of 655.8 nm (Figure 1b). Sensitivity of the detection system can be refined by varying both the driving voltage of the laser and the gain of the PMT. These parameters can be adjusted on a per assay basis to maximize the dynamic range of fluorescence measurements.

Fig. 1.

Fig. 1.

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a) Rendering of the platform illustrating the swiveling of the heater into place over the microfluidic disc which is mounted to an assembly of the motor and optical system. b) Schematic depicting the primary elements of the optical system assembly c) Photograph showing the instrument in the open position. d) Screenshot of the software interface with input parameters (left) and fluorescence readout (right).

The heating system uses a custom medium-wave twin-tube infrared emitter (Heraeus Noblelight) measuring 50 × 22mm. It is mounted in an enclosure that swivels into close proximity to the microfluidic disc and aligns the heating element to a radial strip on one side of the disc. The enclosure is secured with a thumbscrew in both its open and closed positions. When closed, a gasket positioned along the perimeter of the enclosure is compressed to form a tight seal, minimizing the loss of heat and the influence of ambient light during optical detections. Since the heater geometry is not distributed over the entire disc, which would significantly increase cost, the disc rotates at 100 RPM to achieve uniform heating of samples for methods requiring elevated temperatures. At this speed, the disc rotates fast enough for a uniform temperature field (i.e. no hot spot at the location of the heater) without suffering excessive cooling effects due to convective heat transfer to the surrounding air. For example, with a fixed heater output calibrated to achieve 65°C at 100 RPM, a motor speed of 3000 RPM yields a steady state temperature of 50°C with a roughly linear trend for the full range of motor speeds. A gold-coated concave mirror is mounted under the disc, opposite the heater, to redirect stray transmitted radiation back to the disc. For accurate, open-loop temperature control, the heating system was calibrated to correlate heater voltage with sample temperature. A modified motor hub was fabricated with a built-in slip ring to directly probe chamber temperature with a micro-thermocouple while the disc rotated during heating. Steady state temperatures were logged over a range of heater driving voltages, yielding a linear relationship between temperature and voltage in the range of relevant temperatures from 20 to 80°C. Measurements at steady state conditions show temperature stability within a 0.1°C range and day-to-day repeatability within 0.5°C. Uniformity and stability of the heating system are illustrated in Figures S1 and S2, respectively. While the open-loop nature of the system does benefit from stable environmental conditions, most importantly ambient temperature, to maximize accuracy, the temperature requirements of the isothermal amplification performed in this study are relatively forgiving, with 100% polymerase activity reported in the range of 60–70°C. This makes the heating system viable for use in a variety of settings without the need for frequent re-calibration.

These three subsystems are assembled using primarily 3D-printed, heat-resistant glass-filled nylon components (3D Systems), which serve as the heater enclosure as well as modular mounting brackets. These brackets mount and align the brushless motor, optical switch, and optical detection system, and allow for quick-swapping to easily accommodate changing functional requirements such different motor speed capabilities or alternative detection schemes requiring a different configuration of excitation source, filters, lenses, and detector. The mounting brackets are secured to a precision-machined aluminum plate that aligns the subsystems. This assembly is secured to a laser cut panel that is bolted to a frame in the lower compartment of a protective case selected for durability and portability. The case is additionally customized with a cooling fan and vent to keep internal components within recommended operating temperatures. The complete assembly is shown in Fig. 1.

The platform is controlled from a software interface developed in-house using C# (Fig. 1d). This software communicates with 1) an off-the-shelf motor controller (Fauhaber) for rotary control sequences and 2) a low-cost OEM data acquisition unit (National Instruments), which sends and receives digital and analog signals to and from a custom control PCB in order to monitor the optical switch state, power the infrared heater, modulate the laser, and measure voltage from the PMT. A dual output power supply (Delta Electronics) is mounted on the interior and distributes 5VDC and 12VDC to the various components. The maximum power consumption of the instrument does not exceed 30W during operation, making it amenable to battery power and easily compatible with a small generator or modest solar power system. The rear of the case features an IEC AC power jack to plug the instrument into a standard 120VAC outlet and a USB-B jack for connecting to a computer.

2.2. Microfluidic disc

The microfluidic discs used in this work were designed for simplicity, affordability, ease of fabrication, and amenability to mass production. Each disc is composed of three layers: two plastic outer layers sandwiching a double-sided pressure-sensitive adhesive (PSA) layer. The outer layers form the top and bottom of the sample chambers and provide rigidity to the disc. The top layer features through-holes for loading and venting purposes. The bottom layer features etched markings to trigger the optical switch to locate the home position prior to fluorescence measurements. These outer layers are laser cut from 1.5 mm thick clear cast poly methyl methacrylate (PMMA) sheets (McMaster). The middle layer defines the microfluidic features and is laser cut from a sheet of 6 mil (150 µm) thick PSA, which is composed of 2 mil thick mylar with 2 mil thick 300LSE adhesive on both sides (Fralock). Compared to other adhesive tape options, 300LSE adhesive exhibits relatively high adhesion to plastic and is the most appropriate for small bond areas. It provides good chemical and humidity resistance and retains its properties for up to 18 months.

Assembly of the microfluidic disc begins with removing any residues and byproducts from laser cutting from the surfaces of the PMMA layers. Each layer is then treated in a UV-ozone cleaner (Samco) for 2.5 min with an ozone flowrate of 0.75 L/min to remove organic contaminants and alter the surface energy of the parts for greater hydrophilicity to facilitate loading of the disc with aqueous solutions. The layers are then assembled using an alignment jig, which features gage pins corresponding to alignment holes in each layer. Once assembled, the disc is vacuum sealed in a bag using a household vacuum sealing system (FoodSaver) and stored for 24 hours before use. This vacuum sealing method applies uniform pressure to the outer layers during bond build-up of the PSA resulting in a strong, bubble-free bond interface.

Three disc designs were used in this work. For immunoassay-only analyses, an array of 20 chambers (each with a total volume capacity of approximately 10 µL) with tapered geometry is used to localize capture beads into a pellet at the chamber tip for optimal detection. For LAMP-only analyses, a similar array of 20 chambers is used, the only difference being a wide chamber tip geometry to maximize the detection window during fluorescence measurement. For performing simultaneous immunoassays and LAMP reactions, a dual-inlet design is used. This configuration is shown in detail in Fig. 2. The design features a set of nine immunoassay chambers, all connected with a zigzag-style aliquoting channel that was inspired by previously published works (Andersson et al., 2007; Huang et al., 2017; Oh et al., 2016). Along the zigzag channel is an alternating set of capillary valves, which separate the zigzag channel from the chambers, and vents, which promote flow and prevent air entrainment. Once the zigzag channel is filled, the disc is spun at 5000 RPM for 2 sec to equally divide a single sample into 1 µL aliquots and distribute them to the array of chambers. A waste chamber at the end of the aliquoting channel collects any excess sample. In addition to the primary inlet used for aliquoting the sample, secondary inlets for each individual chamber allow the user to load target-specific reagents on a per-chamber basis. Image analysis of scans of the disc at each step of the aliquoting process reveal chamber-to-chamber volume deviation of 1%. A second aliquoting channel connects an array of nine LAMP chambers. A reflective mask, which is laser cut from 1 mm PMMA and covered with adhesive-backed aluminum foil, is secured to the disc to selectively shield the immunoassay chambers from infrared radiation, as illustrated in Fig. 2b. This prevents the immunoassay reagents from reaching excessively high temperatures during the 65°C incubation step for the LAMP reactions. Temperature measurements using an infrared camera (FLIR) indicate that during a 45 min incubation at 65°C, the immunoassay chamber temperatures do not exceed 40°C.

Fig. 2.

Fig. 2.

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a) Schematic of the dual-inlet microfluidic disc indicating the two chamber designs, the zigzag aliquoting channel, and the position of the reflective mask over the immunoassay chambers. b) Exploded view of the mask preventing infrared radiation from overheating the immunoassay chambers while the disc rotates under the heater. c) Scans of the disc during stages of loading using both the chamber-specific inlets and the aliquoting channel.

2.3. Toxin detection via immunoassays

Immunoassays were developed for the detection of three enteric toxins: cholera toxin, SEB, and Stx1. Protocols are similar to previously reported studies (Koh et al., 2015; Phaneuf et al., 2016; Schaff and Sommer, 2011) and summarized here. Cholera antibodies were provided by USDA. Antibodies against SEB and Stx1 were purchased from Toxin Technology. Capture beads are prepared using EDC/NHS chemistry to conjugate 1 µm carboxyl silica microspheres (Bangs Laboratories) with capture antibody at a surface saturation level of 15–20%. Silica beads are incubated with EDC/NHS in MES for 20 min, and then washed with PBS. Activated beads are then conjugated with antibody in PBS with sodium bicarbonate to adjust to pH 8, overnight at room temperature. Conjugated beads are washed in PBS and stored in assay diluent at 4°C until use. To generate the detector, antibodies are labeled with Alexa-647 using the Alexa-647 Antibody Labeling Kit (Life Technologies) according to the kit protocol and incubated in the dark at room temperature for 15 min to yield a label to antibody ratio of 10–15. Instead of using the purification columns provided in the kit, the labeled antibodies are purified using Zeba Spin Desalting Columns (Thermo Fisher Scientific), 7 kDa molecular weight cut-off (MWCO) from Thermo Fisher. The detector antibodies are stored at 4°C until use.

The assay reaction volume is composed of 3 μL capture beads, 1 μL antigen control, and 1 μL detection antibody per chamber. Preparation of this assay begins with combining 15 μL capture beads with 5 μL antigen control (enough for three reactions with excess to offset volume loss) and incubating for 5 minutes. Next, the detection antibody is diluted to 25 nM and 5 μL is added to the capture bead and antigen control solution and allowed to incubate for 20 minutes at room temperature in the dark. For each chamber, 5 μL of this assay mixture is pipetted into a disc preloaded with 1.5 μL density medium per chamber. The disc is sealed with an adhesive backed film (Biorad) then loaded into the instrument. The heater enclosure is swiveled over the disc and secured with its thumbscrew to provide a dark environment for the optical detections. The software is then initiated, performing a pre-centrifugation multipoint scan of the baseline fluorescence of the chambers, spinning at 5000 RPM for 2 min, then performing a second fluorescence measurement of the disc to determine if the toxin was detected. The software reports the median values of each chamber scan to exclude outliers. The PMT voltage values corresponding with fluorescence intensity are reported as relative fluorescence units (RFU). Control experiments using conventional ELISA were performed according to a previously published protocol (Phaneuf et al., 2016). Dose response curves and a method comparison of the microfluidic immunoassays and ELISA can be found in Fig. S3 and S5, respectively.

2.4. Bacteria detection via LAMP

A panel of LAMP reactions was developed for the detection of C. jej, E. coli, and S. typ. Reactions consist of 1 μL of target sample in 9 μL of LAMP reaction mix containing 1× Isothermal Amplification Buffer (20 mM Tris-HCl, 10 mM (NH4)2SO4, 50 mM KCl, 2 mM MgSO4, 0.1% Tween 20, pH 8.8@25°C) (NEB), 800 mM of Betaine (Sigma), 0.14 mM of each nucleotide, 6 mM MgSO4, 0.2 μM each F3 and B3, 0.8 μM each LF and LB, 1.6 μM each FIP and BIP, 1.6 μM quencher primers (Table S1, see Supplementary material), and 3.2 units of Bst 2.0 WarmStart DNA Polymerase (NEB). Primer designs were sourced from published designs (Hara-Kudo et al., 2005; Wang et al., 2012; Yamazaki et al., 2009) and combined with in-house designs for the Cy5 label and addition of a quencher. The use of quencher primers is based on recent work by colleagues towards the enhancement of LAMP chemistry with a technique known as Quenching of Unincorporated Amplification Signal Reporters (QUASR), which is used to increase signal-to-noise for our bacteria detections (Ball et al., 2016). Heat-killed bacterial cells for C. jej, E. coli, and S. typ are sourced from KPL (Cat. No. 50–92-93, 50–95-90, and 50–74-01, respectively). For testing specificity, each reaction is run with a pool of all three bacterial targets and negative template controls were run with a pool of the non-targeted cells. For example, for testing the specificity of C. jej, positive controls contain a mixture of C. jej, E. coli, and S. typ cells, and negative controls contain a mixture of E. coli and S. typ cells. For testing sample matrix independence, a 5% w/v suspension of healthy human fecal matter (Lee BioSolutions, Cat. No. 991–18) is prepared, then spiked with bacterial cells and diluted 1:10 to reduce the concentration of debris. Human sample matrices were acquired from commercial sources from pooled, healthy, adult donors. All biomaterial work was approved by the Institutional Biosafety Committee (project registration number 2018–06-24-CK) and is exempt from IRB review per NIH guidelines. Universal precautions were observed in the handling of all biological materials.

Reactions are performed by first loading 1 μL of the target sample, which is spun into the tip of each chamber. Then 9 μL of LAMP reaction mix is loaded and the inlets of the disc are covered with a sealing film. A reflective mask covering the central surface area of the disc is aligned to expose only the reaction volumes to infrared radiation and the disc is loaded into the instrument. The heater is swiveled into position over the disc and the analysis is initiated through the software interface. After a 10 sec spin at 5000 RPM to ensure complete sample loading and to remove any bubbles, the disc is rotated at 100 RPM for 45 min while the heater is powered at 28 W to maintain 65°C for 45 min. The disc is then spun at 5000 RPM for 5 min to remove bubbles and expedite cooling of the samples via forced convection to <30°C. Finally, each chamber is scanned for fluorescence, just as done for the immunoassays, and the median RFU values are plotted in the software GUI. As a final confirmation step, each disc is subsequently analyzed with a gel imager (Protein Simple). Control experiments using a benchtop thermocycler (Bio-Rad CFX96) were conducted to verify and benchmark the LAMP-based detection of the bacteria panel using the above described protocol for 10 μL reaction volumes. Results are shown in Fig. S4 in the Supplementary material.

2.5. Orthogonal detection via simultaneous immunoassay and LAMP

To perform orthogonal detection, i.e. co-detection of both enteric toxins and bacteria, with minimal steps, reagents for the immunoassays and LAMP reactions are first prepared as described in Sections 2.3 and 2.4. Mock samples are then made by pooling all three toxins for the immunoassays and all three bacterial cells for LAMP. The dual-inlet disc, shown in Fig. 2, is then loaded first with density medium in the tapered immunoassay chambers. Next, toxin-specific capture beads are loaded via the secondary inlets of the immunoassay chambers. Next, the mock samples are loaded into their respective inlets and allowed to flow through the zigzag aliquoting channel and into the waste chamber. A 2 sec spin at 5000 RPM distributes the 1 μL sample volumes into each chamber. Next, toxin-specific detection antibody and bacteria-specific reaction mixes are loaded into their appropriate chambers and another 2 sec spin at 5000 RPM completes the loading process. The mask is then installed and the disc is loaded into the instrument. Following a blank scan of each chamber, the disc is heated to 65°C for 45 min while rotating at 100 RPM to facilitate the LAMP. Next, a high-speed spin at 5000 RPM for 5 min simultaneously cools the disc and performs the sedimentation for the immunoassays. A final scan of fluorescence detects the presence or absence of toxin and bacteria targets, referencing known negative control-based thresholds to verify positive and negative results. Although the incubation step for a standalone immunoassay analysis is typically only 20 min, the extended incubation of 45 min necessitated by the LAMP assays does not adversely affect immunoassay performance.

3. Results and discussion

3.1. Sensitivity

In order to determine the limit of detection of the immunoassays, toxin solutions were diluted to concentrations of 1000, 300, 100, 30, 10, 3 and 0 ng/mL and each was run in triplicate – with the exception of the highest concentration, which was run in duplicate – per disc to demonstrate intra-assay repeatability. Three discs per toxin were analyzed to demonstrate inter-assay repeatability. The resulting dose response curves are shown in Fig. 3.

Fig. 3.

Fig. 3.

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Dose-response curves for cholera toxin, SEB, and Stx1 for the determination of assay sensitivity. Results for each toxin are the combination of three separate disc analyses and are shown as means ± standard deviation, n=9 (except at 1000 ng/mL, n=6). Note: errors smaller than data marker not shown.

Limit of detection values were calculated using a three-parameter fit of log(dose) versus response data. Mean values for each set of three discs for cholera toxin, SEB, and Stx1 were 2.02, 1.35, and 5.50 ng/mL, respectively. This performance is comparable to traditional ELISA, which was used for the same toxin concentrations and yielded limits of detection for cholera toxin, SEB, and Stx1 of 0.33, 2.34, and 1.69 ng/mL, respectively (Fig. S3, see Supplementary material). These results also compare favorable to the lethal dose values for these toxins, reports of which vary depending on the animal model and method of delivery, but can be estimated as follows: 250 μg/kg for cholera toxin, 20 μg/kg for SEB (Gill, 1982), and 20 μg/kg for Stx1 (Tesh et al., 1993). The average inter-assay coefficients of variation (CV) for cholera, SEB, and Stx1 were 13%, 7%, and 16%, respectively. Intra-assay CV values were 10%, 6%, and 12%, respectively. It should be noted that some inter-assay variation can be traced to small changes of the gain values used for the PMT, which was adjusted on a per-assay basis to maximize the dynamic range of each scan.

Similarly, for determining the sensitivity of on-disc LAMP-based detection, serial ten-fold dilutions of bacterial cells were prepared ranging from approximately 103 cells down to single digit cell counts. Each concentration was tested in triplicate. Mean values for each target are plotted in Fig. 4.

Fig. 4.

Fig. 4.

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Plots demonstrate sensitivity of LAMP for detection of the panel of enteric bacteria over a range of starting cell counts. Results are shown as means ± standard deviation, n=3. Red dotted lines represent the smallest measure detected with reasonable certainty.

The limits of detection for C. jej, E. coli, and S. typ are shown to be approximately 5, 1, and 30 cells, respectively. Using the IUPAC definition of the limit of detection, which is the lowest concentration of an analyte determined to be statistically different from an analytical blank, the smallest measure that can be detected with reasonable certainty was calculated from negative control data with 99% confidence level and plotted as dotted red lines for all LAMP data (Long and Winefordner, 1983; McNaught and McNaught, 1997). Although reports on minimum infectious dose vary, often depending on the susceptibility of the population studied, estimates indicate target LOD for viable assays should be on the order of 500 cells of C. jej (Bennett et al., 2014), <700 cells of E. coli O157:H7 (Tuttle et al., 1999), and 10s of cells of S. typ (Kapperud et al., 1990). Since the screening method described here is not targeted at food or environmental sample testing, where infectious dose is a useful parameter, but instead is aimed at detecting the presence of pathogens in biological specimens from already infected patients, bacteria concentrations observed in stool provide a better reference point for performance. For bacteria such as C. jej and S. typ, studies report concentrations ranging from 106 to 109 CFU/g in human stool (Brachman and Abrutyn, 2009). Similarly for E. coli, concentrations have been reported at >106 CFU/g in human stool (Karch et al., 1995).

To confirm that the LAMP assays performed on our microfluidic platform are comparable when performed using a conventional benchtop instrument, serial dilutions of each target bacteria were incubated at 65°C for 45 min, cooled to 25°C, and detected using a Bio-Rad CFX96 (Fig. S4, see Supplementary material). Additional comparisons of these LAMP assays to reference methods can be found in the work done to develop the primer sets used here. The detection of C. jej via LAMP correlated closely with conventional culture-based detection (Yamazaki et al., 2009). The LAMP assay developed for E. coli was shown to be more sensitive than qPCR, especially when working with complex sample matrices such as beef and stool (Wang et al., 2012). Similarly, LAMP assays for the detection of Salmonella serotypes were more sensitive than conventional PCR methods (Hara-Kudo et al., 2005).

3.2. Specificity and sample matrix independence

The specificity and sample matrix independence of the immunoassays have been reported in previous work (Phaneuf et al., 2016) and were shown to be excellent. In order to demonstrate the specificity, specifically exclusivity, of the QUASR-LAMP reactions used in this work, a set of pooled cell mixtures was prepared with final concentrations for each target of 5×105 cells/mL, which is equivalent to 107 cells/g for a 5% w/v stool suspension and therefore at the lower range of concentrations encountered with clinical stool samples. A pool of all target cells was used for the positive controls and pools of non-target cells were used for negative controls. Each was performed in duplicate and can be seen in Fig. 5a. The results indicate clear exclusivity with no false positives, demonstrating that the primer sets and reaction conditions are well-tuned to only the target cells. The need to discern the causative infectious pathogen without non-specific interference is critical, especially considering the large number of endogenous bacteria shed in stool along with the high prevalence of co-infections.

Fig. 5.

Fig. 5.

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a) Specificity of LAMP detection was demonstrated with samples containing a pool of all target cells and negative controls containing only non-target cells. b) Matrix independence of LAMP detection was demonstrated with samples spiked into human stool sample. Red dotted lines represent the smallest measure detected with reasonable certainty.

Sample matrix independence for stool was also tested to demonstrate adequate robustness of the LAMP-based detection. Although stool preparation often requires a series of centrifuging, heating, and pipetting steps (Song et al., 2005), the high sensitivity of our platform allowed for a simple preparation protocol in which a homogenized 5% w/v stool suspension is spiked with bacterial cells at 5×106 cells/mL in a 10:1 volume ratio, yielding a final concentration of 5×105 cells/mL or 107 cells/g, which studies show to be clinically relevant for our panel of bacteria. This solution is then diluted tenfold to reduce stool particulate concentration while maintaining a detectable concentration of 5×104 cells/mL, or 50 cells/reaction. This protocol was used to prepare mock samples for each bacterial target, which were run in duplicate alongside negative controls containing stool only. The results, shown in Fig. 5b, confirm the reputation of LAMP as tolerant to the presence of potential inhibitors relative to other common molecular diagnostic techniques. These results also demonstrate the ability of the platform to tolerate complex clinical samples without a loss in sensitivity, selectivity, or ease-of-use.

3.3. Orthogonal detection of enteric toxin and bacteria panel

Simultaneous immunoassay and LAMP-based detection of both toxins and bacteria was performed using the protocol detailed in Section 2.5. For the immunoassays, a mock sample combining cholera toxin, SEB, and Stx1 was prepared, each with a final concentration of 30 ng/mL, below the thresholds of lethality for each toxin. For the LAMP reactions, C. jej, E. coli, and S. typ were pooled with final concentrations of 5×105 cells/mL (500 cells/reaction). As shown in Fig. 6, all targets were successfully detected on a single disc, yielding signals above the average negative control threshold shown with red dashed lines.

Fig. 6.

Fig. 6.

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Orthogonal detection of bacteria and toxin targets demonstrated for the full enteric panel on a single disc. Red dotted lines represent the smallest measure detected with reasonable certainty.

4. Conclusions

Our focus on the needs of the end user and the realities of point-of-care applications, seeking simplicity without sacrificing efficacy, has yielded a portable, robust screening tool requiring minimal pre-analytic sample handling and preparation. The core feature of the platform that enables multiplexed detections without the burdensome complexity found in similar centrifugal microfluidic devices is the non-contact heating system. The advantages of this system, in which the heat source is spatially separate from the disc, are numerous, including the viability of a simple, low-cost disc, since features such as embedded heating elements or actuation systems for bringing a heater into contact with the disc are not needed. The specific advantage of radiative heating is the ability to introduce optical elements to control temperature by modulating incident radiation with an opaque barrier or neutral density filters for intermediate temperature states. In addition, changing the rotational speed of the disc during heating in order to manipulate convective cooling can also be used to modulate temperature. This heating system, when combined with a precision rotary control system and a high-sensitivity optical detection system, yields a versatile, portable technology capable of providing accurate diagnostic data for myriad purposes, such as guiding the treatment of individual patients, helping combat the increasing levels of antimicrobial resistance, aiding in outbreak tracking and routine public health surveillance, and driving targeted vaccination programs.

Future work will focus on improving portability by integrating a high capacity lithium ion battery to allow for multiple analyses on a single charge. In addition to eliminating the need for an external power source, a built-in touchscreen display and control interface is in development, obviating the need for an external computer. Another planned improvement to the user interface is data interpretation to provide YES/NO detection readouts to the end user. Ease-of-use could be improved by implementing dry storage of assay-specific reagents on-disc, requiring the loading of only the sample and diluent and thereby eliminating multiple preparation steps, saving precious time and reducing opportunities for user error.

Supplementary Material

1
2

Highlights.

  1. Portable, microfluidic platform for detection of proteins and nucleic acids

  2. Simultaneous detection of toxins and pathogenic bacteria from stool

  3. Highly sensitive and specific assays require no sample preparation

  4. Disposable, plastic, microfluidic devices integrate sample handling

  5. Suitable for deployment in low-resource settings

Acknowledgements

This paper describes objective technical results and analysis. Any subjective views or opinions that might be expressed in the paper do not necessarily represent the views of the U.S. Department of Energy or the United States Government. This work was supported by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under award number R01AI098853 and by the Department of Homeland Security, HSARPA-Chemical Biological Division under Agreement HSHQPM-13-X-00222. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or the Department of Homeland Security. Cholera antibodies were produced by Larry Stanker at the United States Department of Agriculture, Agriculture Research Service. Sandia National Laboratories is a multimission laboratory managed and operated by National Technology and Engineering Solutions of Sandia, LLC, a wholly owned subsidiary of Honeywell International, Inc., for the U.S. Department of Energy’s National Nuclear Security Administration under Contract DE-NA0003525. SAND 2018-XXXXJ.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Contributor Information

C. R. Phaneuf, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

B. Mangadu, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

H. M. Tran, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

Y. K. Light, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

A. Sinha, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

F. W. Charbonier, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

T. P. Eckles, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

A. K. Singh, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

C.-Y. Koh, Sandia National Laboratories, 7011 East Avenue, Livermore, CA, 94550

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