BACKGROUND: The wide array of pathogens responsible for infectious diseases makes it difficult to identify causative pathogens with single-plex tests. Although multiplex PCR detects multiple targets, it is restricted to centralized laboratories, which delays test results or makes multiplexing unavailable, depriving healthcare providers of critical, real-time information.
METHODS: To address the need for point-of-care (POC) highly multiplexed tests, we propose the 2-stage, nested-like, rapid (<40 min) isothermal amplification assay, dubbed rapid amplification (RAMP). RAMP's first-stage uses outer loop-mediated isothermal amplification (LAMP) primers to amplify all targets with recombinase polymerase amplification (RPA). First-stage amplicons are aliquoted to second stage reactors, each specialized for a specific target, to undergo LAMP. The assay is implemented in a microfluidic chip. LAMP amplicons are detected in situ with colorimetric dye or with a fluorescent dye and a smartphone.
RESULTS: In experiments on a benchtop and in a microfluidic format, RAMP demonstrated high level of multiplexing (≥16); high sensitivity (i.e., 1 plaque-forming unit of Zika virus) and specificity (no false positives or negatives); speed (<40 min); ease of use; and ability to cope with minimally processed samples.
CONCLUSIONS: RAMP is a hybrid, 2-stage, rapid, and highly sensitive and specific assay with extensive multiplexing capabilities, combining the advantages of RPA and LAMP, while circumventing their respective shortcomings. RAMP can be used in the lab, but one of its distinct advantages is amenability to simple implementation in a microfluidic format for use at the POC, providing healthcare personnel with an inexpensive, highly sensitive tool to detect multiple pathogens in a single sample, on site.
The ability to concurrently detect multiple pathogens infecting a host is crucial for accurate diagnosis of infectious diseases, identification of coinfections, assessment of disease state and treatment efficacy, assessment of drug resistance, and profiling of microbial flora. Pathogens that cause infectious diseases are often coendemic, making it imperative to distinguish single infections from coinfections, which can alter host responses, disease prognosis, transmission dynamics, and treatment strategies and outcomes (1–6). Syndromic panels for infectious diseases can assist healthcare personnel to determine effective treatments and the potential need for additional ancillary diagnostic testing. Standard, culture-based microbiological tests suffer from long turnaround times, ranging from hours to days, and require skill and laboratory facilities. Moreover, culture conditions for many organisms of interest are not known (7). Parasitological and serological methods for detecting infections can be inaccurate, labor-intensive, and unreliable (8, 9).
Molecular methods, particularly PCR, have expanded the range of pathogens that can be identified and greatly shortened time to detection. Existing diagnostic assays are, however, either limited in scope or highly complex (10). Although multiplex PCR (mPCR)3 has the potential to amplify many nucleic acid (NA) targets in a single reaction (11, 12) and, in principle, is suitable for multipathogen detection, it has several limitations. Nonspecific products generated through primer–primer interactions may interfere with the amplification of targets, decreasing sensitivity and selectivity. Moreover, detection of multiple amplicons requires bead arrays, microarrays, or probes with different color fluorophores, which complicates the assay and increases cost (13–15).
Recently, nested mPCR (nmPCR) (16–18) and 2-stage amplification combining PCR and loop-mediated isothermal amplification (LAMP) (isoPCR) (19, 20) have emerged as powerful methods to concurrently detect multiple pathogens. Both involve 2 successive steps of NA amplification, with the first step comprising mPCR and the second step either target-specific PCR or LAMP (21). Amplicons of the first step serve as templates in the second step. Both nmPCR and isoPCR significantly enhance limits of detection and specificity over single-stage, mPCR. However, both methods require at least one thermal cycling (PCR) process. Thermal cycling complicates instrumentation and is not compatible with point-of-care (POC) applications. Moreover, the need to transfer first-stage amplicons to the second-stage exposes the NA-rich first-stage tube to the environment, potentially contaminating the work space. A closed system for 2-stage amplification is, therefore, preferable.
In recent years, there has been a growing interest in isothermal amplification methods (22) such as recombinase polymerase amplification (RPA) (23) and LAMP (21) for POC diagnostics. A few groups have developed multiplexed RPA and multiplexed LAMP assays (24–28) for codetection of a small number (i.e., ≤4) of targets (24–28).
To improve the sensitivity of the amplification process, enable concurrent detection of multiple NA targets, and benefit from the advantages of isothermal amplification (such as simple instrumentation and low power consumption), we propose a 2-stage, isothermal–isothermal, enzymatic amplification method, dubbed rapid amplification (RAMP), that consists of a first-stage RPA and second-stage LAMP. To demonstrate RAMP's capabilities, we first carried out benchtop experiments to optimize the assay and compare its sensitivity to that of LAMP alone and RPA alone. Second, we examined multiplex assays to detect pathogens that are prevalent in low-resource settings, and, in some cases, coendemic. To eliminate the need for pipetting, to carry out the entire RAMP assay in a closed system, and to enable testing by minimally trained personnel, we implemented the RAMP assay in a microfluidic format.
RAMP consists of 2 successive isothermal enzymatic amplification stages. The first stage is a RPA isothermal amplification at approximately 37 °C for 10–20 min, with primers for all the targets. First-stage amplicons are then distributed among individual LAMP reactors, each specific to a target, and serve as templates in second-stage LAMP reactions at 60–65 °C, typically for 30 min. LAMP is monitored in real time, with either a nonspecific intercalating fluorescent dye or a colorimetric dye.
The first-stage RPA uses a mixture of all F3 and B3 primers of the second LAMP stage concurrently amplifying all the targets present in the sample (Fig. 1). In contrast, each of the second-stage reactors operates with a set of 6 LAMP primers and amplifies a specific target. Typical amplicons' length range from 180–330 bp. RAMP was implemented either on benchtop (Fig. 1A) or in a microfluidic format with spontaneous distribution of first-stage products to second-stage reactors (Fig. 1C).
In our experiments, we used different types of samples: (a) purified DNA and RNA, (b) serum from Schistosoma mansoni-infected mice, (c) spiked human serum and whole blood, and (d) spiked urine samples. See Supplemental Materials and Methods in the Data Supplement that accompanies the online version of this article at http://www.clinchem.org/content/vol63/issue3 for additional details of the experimental procedures.
We first describe our benchtop experiments with the RAMP assay, which, among other things, serve to optimize the assay. Then, we describe the implementation of the assay in a microfluidic format.
We first investigated RAMP's performance with purified samples with single targets on the benchtop (see Supplemental Results 1 in the online Data Supplement). RAMP was consistently more sensitive (typically, ×10) than LAMP alone and more specific and efficient than RPA alone. Any spurious first-stage RPA amplicons did not amplify in the specific second-stage LAMP and did not produce any detectable signals. In contrast to RPA (23), RAMP's amplicons can be detected in situ with nonspecific dyes without a need to discharge the amplicons to a hybridization array.
Next, we examined the effects of first-stage (RPA) reaction time and primer combinations on overall RAMP performance (see Supplemental Results 2 in the online Data Supplement). A 20-min RPA, followed with a 15–20 min LAMP yielded detectable signals even in the presence of low-abundance targets. Specifically, the single-plex RAMP detected successfully 0.5 fg S. mansoni DNA. We also examined various primer combinations. RAMP assay with F3-B3 primers in stage 1 and the 4 primers FIP, BIP, Loop F, and Loop B in stage 2 provided highly specific results and were used throughout this report. The sequences of the various primers used in this work are listed in Supplemental Table 1A in the online Data Supplement.
MULTIPLEX RAMP ASSAY
Although RAMP has higher sensitivity than LAMP and is useful for this reason alone, RAMP's main benefit is its capacity to detect multiple targets. We first carried out a 4-plex RAMP assay for the detection of HIV, S. mansoni, Plasmodium falciparum, and Schistosoma hematobium, and found it highly sensitive and specific (see Supplemental Results 3 in the online Data Supplement). Encouraged by this successful performance, we increased the level of multiplexing to 16 targets. We designed an assay to detect S. mansoni, HIV-1 clade B, S. hematobium, P. falciparum, Schistosoma japonicum, Brugia malayi, Strongyloides stercoralis, drug-resistant Salmonella, Zika virus (ZIKV)-America strain (mex 2–81, Mexico), ZIKV-Africa strain (MR 766, Uganda), human papilloma virus (HPV)-58, HPV-52, HPV-35, HPV-45, HPV-18, and HPV-16. The primers' sequences are listed in Supplemental Table 1A in the online Data Supplement. The above targets were selected for proof of concept, taking advantage of reagents and targets available in our lab, and not for clinical reasons (though many of the selected pathogens are coendemic). The targets are both DNA and RNA (HIV-1 and ZIKV), and range from viruses to multicellular metazoans.
Fig. 2 depicts amplification curves of samples containing the following according to the panels listed here: (A) HPV-16 (100 copies) and ZIKV [50 plaque-forming units (PFU), American strain]; (B) HPV-18 (100 copies) and ZIKV (50 PFU, African strain); (C) HIV-1 clade B (100 copies), P. falciparum (300 fg DNA), the schistosome S. japonicum (1 pg DNA), the filarial nematode B. malayi (13 pg DNA), the soil-transmitted nematode S. stercoralis (1 pg DNA), and drug-resistant Salmonella (100 copies); and (D) no targets (negative control). Once again, RAMP proved to be highly sensitive and specific, with no false positives or negatives. At the tested concentrations, the assay successfully discriminated among various strains of HPV and between American and African strains of ZIKV.
To examine assay sensitivity and the dependence of the threshold time on target concentration, we repeated our experiments by using a dilution series. In the interest of brevity, we show results only for the American strain of the ZIKV. Fig. 2E depicts the amplification curves obtained with the 16-plex RAMP assay in the presence of 0, 1, 5, 50, and 500 PFU of the American ZIKV. Note that 1 PFU of ZIKV is detected. When the number of Zika templates was equal to or larger than 5 PFU, the threshold time [the time required for the signal to reach half its saturation value (T1/2)] was a linear function of the number of target ZIKV (PFU) (Fig. 2F) and data were highly reproducible (with a relative standard deviation in threshold time of 2%). If we were to process a 100-μL sample, the RAMP's detection limit for ZIKV would be better than 50 PFU/mL. This is orders of magnitude lower than the ZIKV concentrations, ranging from 103–106 PFU/mL, in symptomatic Zika-infected patients (29). Moreover, our data suggest that multiplex RAMP can genotype HPV strains and, at the target concentrations tested, differentiate the American ZIKV from the African strain.
To reduce test complexity, it is occasionally desirable to minimize, or even eliminate, sample preparation. To assess RAMP compatibility with minimally processed samples, we used RAMP to detect pathogens in urine, serum, and whole blood samples without NA extraction (see Supplemental Results 4 in the online Data Supplement). RAMP demonstrated robustness and was refractory to inhibitors. Specifically, the16-plex RAMP of this section successfully and reproducibly detected 5 PFU ZIKV when 5 μL urine was added directly to the first-stage reaction volume.
IMPLEMENTATION OF RAMP IN A MICROFLUIDIC FORMAT
To enable the use of RAMP at the POC by minimally trained personnel, avoid the tedium of pipetting first-stage products into multiple second-stage reactors, and minimize risk of contamination, it is desirable to automate the process of distributing first-stage products into second-stage reactors and implement it in a closed system. Fortunately, these objectives can readily be achieved.
We designed, fabricated, and tested a prototype of a microfluidic chip for multiplex detection of DNA and RNA targets with RAMP. Fig. 3 and Fig. 1C show, respectively, 4-plex and 16-plex chips. Fig. 3A depicts schematically the structure of the 4-plex plastic chip, which includes a main chamber for the first-stage RPA reaction and 4 branching chambers for second-stage LAMP amplifications (the number of these branching chambers can be adjusted according to the number of desired targets as shown in Fig. 1C). In the embodiment shown in Fig. 3A, the first-stage 25-μL (RPA) amplification chamber is in direct communication with the second-stage chambers, each 15 μL in volume. In another embodiment (with which we have experimented, but do not discuss here), we sealed the connectors between the first-stage chamber and the second-stage chambers with paraffin that melts once the chip's temperature exceeds 60 °C. During the first stage (37 °C), the paraffin remains solid and blocks the passages between the first-stage RPA chamber and the LAMP chambers. When the chip's temperature increases to 65 °C for the second-stage LAMP, the paraffin melts, opening the passages and allowing the RPA products diffuse into the second-stage LAMP chambers to serve as templates in the second-stage amplification.
Our hypothesis is that the first-stage amplicons are uniformly distributed in the first-stage chamber. This hypothesis is borne out by the reproducibility of our data, but requires further study. The rate of diffusion of the amplicons from the first stage to the second stage is controllable by the size of the passages leading from the first-stage chamber to second-stage chambers.
In the experiments described here, the targets and RPA cocktail were introduced into the first-stage chamber and incubated at 37 °C for a predetermined time (15 min). The first-stage amplicons diffused into the second-stage chambers. After 15 min, the chip was heated to 63 °C to carry out LAMP amplifications. The prestored LAMP reagents (currently, in liquid form) included either an intercalating fluorescent dye such as EvaGreen or a colorimetric dye such as Leuco crystal violet (LCV) (30). The fluorescence emission and the change of color in the second-stage chambers were, respectively, monitored during the amplification process with a smartphone camera (31) and by eye. We did not observe any crosstalk among the second-stage chambers. Nevertheless, in future implementations, we will encapsulate the lyophilized LAMP reagents in paraffin for refrigeration-free, long shelf life and just-in-time release (32).
As a proof of concept, we prestored in the second-stage chambers LAMP primers targeting HIV clade B (chambers 1 and 3), drug-resistant Salmonella (chamber 2), and S. mansoni DNA (chamber 4). We then added 1000 copies of HIV RNA and 100 copies of drug-resistant Salmonella genomic DNA (gDNA) (and no S. mansoni gDNA) in RPA reaction buffer and inserted the reaction buffer in the first-stage chamber. We monitored the emissions from the LAMP reactors as functions of time (see the Supplemental Video in the online Data Supplement). Fig. 3B depicts the fluorescence emission from the second-stage reactors 50 min after the start of the assay. The second-stage reactors specific for HIV (2 reactors) and Salmonella emit strong fluorescence signals. The S. mansoni reactor that serves here as a negative (no target) control emits no signal, as expected. This experiment indicates that the RAMP assay is amenable to simple microfluidic implementation that eliminates the need for manual transfer of first-stage amplicons to the second stage. Moreover, the transfer from the first to the second stage takes place in a closed system, eliminating the risk of contaminating the environment with NAs or picking up contaminants from the environment.
Although we can excite fluorescence with a cell phone flash and detect emission with the cell camera (31), this requires the use of filters to separate between the excitation and emission spectra, which may add slightly to the device's cost. To further simplify the assay, we replaced fluorescence-based detection with a colorimetric LCV (30) dye, allowing us to read the RAMP results by naked eye, without any detector, or monitor them with a cell phone camera.
Fig. 3C shows colorimetric detection of 100 copies of HIV RNA targets in PCR vials. Notice the well-defined color contrast between the colorless negative control and the violet positive test. The dye-based detection can be readily implemented into our microfluidic format. Fig. 3D depicts examples of microfluidic RAMP assays with different samples, by using dye-based detection. The second-stage reactors were specialized to detect HIV-1 clade B, S. mansoni, P. falciparum, and S. hematobium. Fig. 3D(i) shows results obtained with a negative (no targets) sample. None of the second-stage reactors turned violet, consistent with negative tests. Fig. 3D(ii) shows results for a sample containing 300 fg P. falciparum DNA and no other targets. Note that only the reactor specific for P. falciparum turned violet, while the 3 other chambers remained colorless, consistent with a positive test for P. falciparum and negative tests for the other 3 targets. Fig. 3D(iii) shows results for a sample containing 300 fg P. falciparum DNA, 1000 copies of HIV RNA, and no other targets. Only the reactors specialized for P. falciparum and HIV turned violet, consistent with a positive test for P. falciparum and HIV and negative for the other 2 targets included in the assay. The multiplexed assay operated well, producing no false positives and no false negatives. To eliminate subjectivity, we found we could delegate the test reading to a smartphone that imaged the signal, compared it to a control, and applied preprogrammed thresholds to discriminate between positive and negative signals.
Different infections can cause similar overt symptoms, but require diverse disease management strategies (i.e., the current ZIKV epidemic) (1, 6). Patients may be subject to coinfections that alter symptoms, immune responses, and therapies (2–5). These are just 2 examples in which multiplexed platforms would provide timely, cost-effective diagnosis. Although PCR-based multiplexed platforms are available, for the most part, they require expensive equipment and highly skilled personnel, which precludes their use in resource-poor settings and causes a significant delay between sample collection and test results. There is a great need in both developed and developing countries to enable healthcare providers and epidemiologists to make timely, informed decisions regarding treatment options and efficacy. Economic pressures also motivate efforts to develop multiplexed diagnostic procedures that can be carried out at the patient's side.
To address this need, we propose a 2-stage, hybrid isothermal enzymatic amplification method, dubbed RAMP, comprising a first-stage RPA reaction, in which all the targets in the sample are amplified concurrently, combined with a second, highly specific LAMP. RAMP merges the advantages of RPA and LAMP, while circumventing their shortcomings. RAMP benefits from RPA's high tolerance to inhibitors while overcoming RPA's tendency to produce spurious amplicons. RAMP can concurrently detect multiple DNA and RNA targets without undue demands on sample volume, operating with volumes similar to those used in single-plex detections.
Specifically, RAMP's first stage uses outer LAMP primers F3 and B3 of all targets while the second stage uses the other 4 RAMP primers. This nested-like principle provides high sensitivity, specificity, and robust, extensive multiplexing capabilities. RAMP's sensitivity is, respectively, about 10- and 100-fold better than LAMP when operating with purified and crude samples. RAMP is also more efficient and specific than RPA. Although spurious amplicons are produced in RAMP's first stage, they do not undergo further amplification in the second stage and their concentration is sufficiently low as not to produce false signal, permitting the use of inexpensive nonspecific dyes to identify the presence of double-stranded DNA (dsDNA).
We demonstrated 2 detection modalities. In a few of our experiments, we used an intercalating fluorescent dye that can be excited with a smartphone flash and detected with a phone camera (31). In other experiments, we adapted the colorimetric dye LCV, which does not require excitation. The color change induced by the presence of amplicons can be visualized by eye or detected with a cell phone camera. Both labeling methods are compatible with POC applications. In contrast, detection of RPA products requires specific molecular beacons and/or hybridization arrays.
Although we applied a 16-plex RAMP to samples that included similar targets, such as different strains of HPV and the ZIKV, the assay did not produce any false positives or false negatives and at the concentration tested was able to discriminate among the various strains. Although more extensive testing is needed, the initial results are highly promising. We demonstrated here a successful 16-plex RAMP. It is likely that concurrent detection of more than 16 targets is feasible.
RAMP has many advantages over conventional multiplexing strategies such as mPCR, nmPCR, and isoPCR. RAMP is faster and more inhibitor-tolerant than mPCR, nmPCR, and isoPCR. RAMP takes <40 min, even in the presence of low-abundance samples. In contrast, amplification times of nmPCR (16, 19, 33, 34) and isoPCR (19) exceed 2 h and 1 h, respectively. Since in our RAMP experiments, we did not optimize the primers, we allocated 15–20 min to the first-stage RPA and 20 min to the second-stage LAMP. With optimized primers, such as elongated F1-B1 primers, it may be possible to further reduce the duration of the RPA (23) and thereby, of the entire RAMP.
RAMP inherits RPA's tolerance to impurities. In the presence of high-abundance targets, RAMP can detect targets in minimally purified samples, which is not possible with PCR (35, 36). By equipping the first-stage reaction chamber with an NA isolation membrane, as we have previously described (37), we decouple the sample volume from the reaction volume, purify the NAs in situ, and achieve sensitivities on par with state-of-the-art laboratory-based equipment. Finally, since RAMP is based on 2 isothermal processes and does not require thermal cycling, it can operate with much simpler instrumentation than PCR. Table 1 provides a qualitative comparison of RAMP with other multiplexing strategies.
One of RAMP's major advantages is its easy implementation in a microfluidic format, wherein first-stage amplicons diffuse into the second-stage reactors without a need for mechanical aliquoting. The entire RAMP process can be carried out in a closed system without exposing the NA-rich, first-stage products to the environment, minimizing contamination risk. The ability to implement RAMP in a simple microfluidic format and its minimal demands on instrumentation, make it an ideal candidate for POC applications: at the sample collection site, next to the patient, at home, in the clinic, or in the field. For POC applications, it is necessary, however, to prestore all reagents in the diagnostic cassette, refrigeration-free. Although we did not discuss reagent storage in this report, we describe a possible strategy elsewhere (32).
In summary, RAMP is a hybrid, 2-stage, rapid, high-sensitivity and -specificity assay with extensive multiplexing capabilities. RAMP can be used in the lab, but one of its distinct advantages is its amenability to implementation at the POC, providing healthcare personnel with a tool to detect multiple pathogens in a single sample without a need to send the sample to a centralized laboratory. We demonstrate the feasibility of the RAMP assay, showing concurrent detection of DNA and RNA from several pathogens, including retroviruses, flaviviruses, papillomavirus, bacteria, protozoa, and parasitic helminths. In the future, we hope to use the RAMP concept to develop various diagnostic panels of medical significance such as vector-transmitted pathogens and HPV genotyping.
↵3 Nonstandard abbreviations:
- multiplex PCR;
- nucleic acid;
- nested mPCR;
- loop-mediated isothermal amplification;
- 2-stage amplification combining PCR and LAMP;
- recombinase polymerase amplification;
- multiplex RPA;
- rapid amplification;
- Zika virus;
- human papilloma virus;
- plaque-forming units;
- the threshold time (the time required for the signal to reach half its saturation value);
- Leuco crystal violet;
- genomic DNA;
- double-stranded DNA.
Author Contributions: All authors confirmed they have contributed to the intellectual content of this paper and have met the following 3 requirements: (a) significant contributions to the conception and design, acquisition of data, or analysis and interpretation of data; (b) drafting or revising the article for intellectual content; and (c) final approval of the published article.
Authors' Disclosures or Potential Conflicts of Interest: Upon manuscript submission, all authors completed the author disclosure form. Disclosures and/or potential conflicts of interest:
Employment or Leadership: None declared.
Consultant or Advisory Role: None declared.
Stock Ownership: None declared.
Honoraria: None declared.
Research Funding: J. Lok, NIH; R. Greenberg, NIH; H. Bau, NIH grant 1R41AI104418-01A1 (to the institution).
Expert Testimony: None declared.
Patents: H. Bau, US provisional patent application no. 62/278,095.
Role of Sponsor: The funding organizations played no role in the design of study, choice of enrolled patients, review and interpretation of data, and final approval of manuscript.
- Received for publication July 13, 2016.
- Accepted for publication October 31, 2016.
- © 2016 American Association for Clinical Chemistry