Recombinase Polymerase Amplification for Rapid Detection of Zoonotic Pathogens: An Overview

Abstract

With the advent of molecular technology, several isothermal techniques for rapid detection of zoonotic pathogens have been developed. Among them, recombinase polymerase amplification (RPA) is becoming an important technology for rapid, sensitive, and economical detection of zoonotic pathogens. RPA technology has the advantage of being able to be implemented in field settings, because the method requires minimal sample preparation and is performed at a constant low temperature (37–42°C). RPA is rapidly becoming a promising tool for the rapid detection, prevention, and control of zoonotic diseases. This article discusses the principles of RPA technology and its derivatives, including RPA coupled with lateral flow testing (RPA-LF), real-time fluorescence RPA, electrochemical RPA, and flocculation RPA, and their applications in the detection of zoonotic pathogens.

Main article text

INTRODUCTION

Nucleic acid amplification (NAA) in vitro, the artificial replication of genetic material, has permeated nearly every field of the life sciences and biotechnology. This technology originated from the invention of polymerase chain reaction (PCR) by Kary Mullis in 1983 [1]. The PCR amplification technique has been widely used in the rapid detection of nucleic acids and demonstrated to be highly specific and efficient. However, the PCR amplification technique has several limitations. First, it is somewhat limited by its reliance on a thermal cycler for heating and cooling, and high-quality nucleic acid reactants. Furthermore, skilled operators and a laboratory environment are also needed, thus preventing its application in low-resource settings.

To overcome the limitations of PCR methods, multiple isothermal DNA amplification methods using various enzymes and amplification systems have been established, including loop-mediated isothermal amplification (LAMP), rolling circle amplification (RCA), nucleic acid sequence-based amplification (NASBA), helicase-dependent amplification (HDA), strand displacement amplification (SDA) and recombinase polymerase amplification (RPA) (Table 1) [2–4]. Isothermal NAA greatly simplifies the incubation conditions for artificial NAA, and the elimination of thermal cycling decreases the need for amplification equipment or a laboratory environment. Another advantage of the isothermal NAA is that multiple molecular reactions, such as denaturation, annealing, and elongation, can be performed asynchronously in one isothermal amplification reaction, thus effectively decreasing the NAA reaction time [5,6]. Among all isothermal DNA amplification methods, RPA is notable for its simplicity, and high specificity and sensitivity, despite its very short history of use.

TABLE 1 |

Isothermal nucleic acid amplification techniques.

Isothermal technique	Template	Primers	Temperature (°C)	Incubation time (min)	Lyophilized reagents
LAMP	DNA	4∼6	60∼65	15∼60	No
RCA	DNA/RNA	1	30∼65	60∼240	No
NASBA	RNA	2	37∼42	60∼180	Yes
HDA	DNA	2	60∼65	30∼120	No
SDA	DNA	4	30∼55	60∼120	No
RPA	DNA/RNA	2	37∼42	20∼40	Yes

RPA

RPA was first developed by Niall Armes at ASM Scientific Ltd of Cambridge, UK in 2006 [7]. Unlike traditional PCR methods, RPA does not rely on thermal denaturation and annealing. Three key proteins required for RPA reactions are a recombinase that binds single-stranded nucleic acid, a single-stranded DNA binding protein, and strand-displacement DNA polymerase [7,8]. The amplification reaction starts with the binding of the recombinase to a primer of approximately 30–35 nt, and the complex’s subsequent search for the target site in the double-stranded DNA template. After the site is located, the complex directly initiates a chain exchange reaction to form a D-shaped loop. Single-stranded binding protein then binds the displaced DNA chain to prevent primer dissociation. Subsequently, the active hydrolysis of ATP by the recombinase-primer complex alters the conformation of the complex. After the recombinase releases from the nucleoprotein filament, the 3′ end of the primer is exposed and recognized by the strand-displacement DNA polymerase. The DNA polymerase adds the corresponding base to the 3′ end of the primer according to the template sequence, and the DNA amplification reaction begins. Both forward and reverse primers allow the amplification reaction to occur in both directions simultaneously. The synthesized amplicon can be used as a new template to achieve exponential amplification Figure 1 [9].

FIGURE 1 |

Schematic of the RPA principle. (A) The recombinase proteins bind each primer. (B) The complex searches for the target site and directly initiates a chain exchange reaction, forming a D-loop. (C) Single stranded binding proteins bind the displaced DNA chain, thus preventing primer dissociation. (D) The recombinase releases the nucleoprotein filament, and the DNA polymerase extends the primer 3´ ends. (E) The DNA amplification reaction starts. (F) Exponential amplification is achieved by cyclically repeating the process.

Template

RPA was initially designed to amplify double-stranded DNA, single-stranded DNA, and DNA methylation [10,11]. The cDNA produced by reverse transcription of RNA or miRNA can also be amplified [12,13] in a process requiring reverse transcriptase [14]. Regardless of the nucleic acid template type, the length of the RPA amplicon should be less than 500 bp for efficient amplification. Most published articles on RPA have shown that although targets up to 1.5 kb can be amplified, this method is more suitable for amplicons of between 100 and 200 bp [7,15], because shorter the sequences result in higher amplification efficiency.

Primers and probes

In contrast to traditional PCR primers, RPA primers are generally 30–35 nucleotides long, to facilitate the formation of a complex between the recombinase and the primer. Longer primers (up to 45 nucleotides) may produce secondary structure and potential primer artifacts, thus decreasing amplification efficiency. In addition, no melting temperature requirement exists for RPA primer and probe design. Like traditional PCR primers, RPA primers should have a GC content of 30%–70%, and long-chain guanines should be avoided at the 5′ end, whereas guanine and cytosine nucleosides can be used at the 3′ end to improve performance. A probe is not necessary in standard RPA assays but is required when RPA is combined with the various endpoint detection methods described below [16–18]. The procedure for primer and probe design is not standardized, and no dedicated software is available; however, the selected primers and probes can be evaluated with software for PCR primer design, such as Primer Premier 5. Multiple groups of primers and probes are usually designed and evaluated in experiments to screen for the best combinations [19–22].

Temperature and incubation time

The optimum reaction temperature of the enzymes in RPA assays is 25–42°C; thus, the optimum temperature for the RPA reaction is also in this range. RPA assays do not require strict temperature control [23–25]. Even if the temperature exceeds the recommended range, the RPA reaction can still proceed. However, the recommended RPA reaction temperature in most studies is 37–42°C [9,26].

The time required for NAA to reach a detectable level depends on the concentration of the starting DNA template. At the appropriate reaction temperature, detection can be completed within 20 minutes [27]. In practical applications, the amplification results can be observed in as few as 3–4 minutes. For solution phase RPA amplification reactions, the recombinase can consume all ATP in the system within 25 minutes; therefore, long incubation times are unnecessary. In addition, a shaking step at the fourth minute of the reaction is recommended to increase reaction efficiency [20,21,28].

Advantages and disadvantages of RPA assays

RPA assays offer several advantages. NAA can be performed under constant temperature conditions of 37–42°C without pre-denaturation steps and high temperature annealing steps. Therefore, RPA does not require expensive thermal cycling equipment and is suitable for non-instrumented NAA platforms. Researchers have also used human body temperature to complete the amplification in various conditions [28].

RPA technology is simple to perform and has good expansibility, and can be combined with various systems for different detection purposes. Combining this method with reverse transcriptase in reverse transcription-RPA (RT-RPA) can be used to amplify RNA sequences; combining RPA with a fluorescence probe in real-time fluorescence RPA can achieve real-time detection; and combining RPA with lateral flow (LF) in RPA-LF can detect enable visual target sequence detection with the naked eye [18,29–32]. Recently, clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated nuclease (Cas) have been used to develop a CRISPR-based diagnostic method called SHERLOCK, thus increasing the popularity and scope of application of RPA assays [33].

RPA reagents can also be used in lyophilized forms with excellent stability, and be transported and stored without refrigeration [34,35]. Multiplex RPA assays that can detect multiple target sequences quickly in a single reaction are also available, depending on the target sequence, amplicon size, and primer design [36–38]. All these advantages facilitate the implementation of RPA in field-based rapid detection applications. In recent years, RPA methods have been extensively developed for the rapid detection of various pathogens, particularly zoonotic pathogens.

Some shortcomings of RPA technology include that RPA products normally require purification before agarose gel electrophoresis to avoid smearing due to the presence of other components. Second, no specialized software is available for the design and screening of RPA primers and probes. Thus, costly and time-consuming synthesis and screening studies are required. Furthermore, conventional real-time PCR probes (such as Taq-Man probes) are not compatible with the RPA reaction, and fluorescent dyes are prone to false positive results. In addition, real-time amplification with RPA is not easily controlled, owing to its isothermal amplification properties and the use of a time threshold rather than a cycling threshold. Thus, the reactions are dependent on the initial conditions, incubation temperature, and mixing steps. With regard to cost, RPA kits are currently sold by only one company, and users have limited flexibility in the kit formulations; consequently, small batch use has high costs.

APPLICATIONS OF RPA IN ZOONOTIC PATHOGEN DETECTION

Zoonoses are any diseases or infections that are naturally transmissible from vertebrate animals to humans, or from humans to animals [35,39]. Approximately 60% of emerging human infections are zoonoses, and more than 70% of pathogens originate from wild animal species [40]. At present, more than 200 species of zoonotic pathogens are known worldwide, the most prevalent of which are anthrax, plague, foot-and-mouth disease, avian influenza, Japanese encephalitis, and rabies [41–43]. Zoonoses are a major public health concern and directly threaten human health. In recent decades, large-scale epidemics of zoonoses have occurred, such as the 2005 H5/N1 avian influenza outbreak, the 2009 H1/N1 influenza pandemic, the 2013–2016 West African Ebola outbreak, and the COVID-19 pandemic [44]. Since the start of the 21st century, the global economic costs of zoonosis outbreaks have exceeded 100 billion U.S. dollars [45,46]. The current COVID-19 pandemic, affecting millions of people worldwide, has stimulated the development of rapid and sensitive technologies for zoonotic pathogen detection, general patient and health care, and prevention of large-scale outbreaks and further spread, which are of paramount importance in public health [47]. RPA assays are widely used in the detection of zoonotic pathogens because of their high sensitivity, efficiency, expansibility, rapidity, and specificity (Table 2) [48–58].

TABLE 2 |

Major zoonotic diseases and RPA detection methods.

Disease	Etiology	RPA method	Amplification time (min)	Temperature (°C)	LOD
Bacterial zoonoses
Tuberculosis	Mycobacterium bovis, Mycobacterium caprae, Mycobacterium microti	Direct RPA	20	39	6.25 fg
		LF-RPA	25–45	5	5 copies/reaction
		Real-time fluorescence RPA	20	39	4 copies/μl
		Electrochemical RPA	20	39	0.04 ng/μl
		CRISPR/Cas-RPA	180	37	4.48 fmol/L
Brucellosis	Brucella abortus Brucella melitensis, Brucella suis, Brucella canis,	Direct RPA	20	38	3 copies/reaction
		LF-RPA	10–30	30–37	6 copies/reaction
		Real-time fluorescence RPA	16	40	17 copies/reaction
Plague	Yersinia pestis	CRISPR/Cas-LF-RPA	50	37	103–106 fg/μl
Leptospirosis	Leptospira interrogans	CRISPR/Cas-RPA	60	39	100 copies/ml
Tularemia	Francisella tularensis	Real-time fluorescence RPA	20	39–42	10 copies/reaction
		Electrochemical RPA	60	37	500 fM
Lyme disease	Borrelia burgdorferi	LF-RPA	30	37	25 copies/reaction
Viral zoonoses
Rabies	Rabies virus	Direct RPA	20	42	562 copies/reaction
		Real-time fluorescence RPA	15	42	4 copies/reaction
Avian influenza	Influenza A virus	LF-RPA	20	30–42	0.15 pg
		Real-time fluorescence RPA	20	39	100 copies/reaction
Ebola disease	Ebola virus	LF-RPA	40	37	134 copies/μl
Dengue fever	Dengue virus	LF-RPA	23	37	10 copies/μl
		Real-time fluorescence RPA	20	38	14–241 copies/reaction
Zika fever	Zika virus	Real-time fluorescence RPA	20	41	5 copies/reaction
West Nile fever	West Nile virus	Real-time fluorescence RPA	15	39	10 copies/reaction
SARS	SARS coronavirus	LF-RPA	45	42	35.4 copies/μl
		Real-time fluorescence RPA	20	42	7.74 copies/reaction
		CRISPR/Cas-RPA	50	37	1–10 copies/reaction
		Ligation-RPA	30	37	10 copies/reaction
		RPA/rkDNA-graphene oxide probing	96	37	6.0 aM

RPA-LF

Most end-point RPA detection methods reported to date rely on LF assays, and the results can be obtained extremely rapidly in a visual read-out format. LF chromatography test strips are mainly used as simple devices for qualitative and semi-quantitative detection, and can be used in resource-limited or non-laboratory environments [59]. RPA-LF is based on the principle of RPA amplification with biotin-labeled primers and carboxyfluorescein (FAM)-labeled probes for amplification reactions with target nucleic acids; the final amplified product carries both FAM and biotin labels. The detection line of the LF test strip contains streptavidin. When the FAM on the amplicon binds the gold-labeled-anti-FAM antibody in the sample pad, an immune complex is formed. The immune complex undergoes chromatographic diffusion on the strip. The streptavidin on the detection line captures the immune complex containing the biotin amplicon, thus resulting in color development [60]. In addition, multiplex LF strips have been developed, such as the PCRD nucleic acid detector cassette (Abingdon Health, UK), which has two detection lines that can detect FAM/biotin and DIG/biotin labeled amplicons, respectively. This method allows for the detection of various pathogens in the same tube as well as the introduction of an internal control, as has been applied in the detection of three Anaplasma species [61].

The limit of detection (LOD) of RPA-LF can be as low as one to ten copies per reaction in the detection of zoonotic pathogens. Wu et al. have established RPA-LF for detecting Toxoplasma gondii, with a LOD of 0.1 oocyst/reaction, a value ten times higher than the sensitivity of nested PCR [62]. Shi et al. have used this method to detect avian influenza A virus (H7N9) with a LOD of 32 fg nucleic acid sample, and without cross-reaction with other subtypes of influenza viruses [55]. Rani et al. have presented a rapid, sensitive, specific, and portable method to detect the rfbE, fliC, and stx genes of Escherichia coli O157:H7, with LODs as low as 4–5 CFU/mL, 10¹ CFU/mL, and 10² CFU/mL, respectively, in 8 minutes at temperatures between 37 and 42°C [53].

RPA-LF has also been used in the detection of other zoonotic parasites, bacteria, and viruses, such as Trypanosoma cruzi, Brucella spp., Burkholderia mallei, Chlamydia trachomatis, Orientia tsutsugamushi, Rickettsia typhi, Coxiella burnetii, Borrelia burgdorferi, Newcastle disease virus, dengue virus, orf virus, human adenovirus, and severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) [19,20,22,28,49,52,53,57,63–71].

RPA-LF detection can be performed in approximately 20 minutes at 25–45°C. Therefore, simple heating equipment, such as electric water heaters, or even human body heat can be used to achieve accurate detection. The RPA-LF test results show a red band on the strip, which can be observed with the naked eye. Even non-professionals can directly observe the analytic results. This method is highly suitable for on-site detection, particularly in areas with poor economic conditions and limited resources [49,72].

However, LF assays’ insufficient accuracy and stability make them unsuitable for quantitative analysis in clinical applications [73]. In addition, during the color development of strips, the lid of the reaction tube must be kept open, thus making the method prone to environmental contamination and false positives Table 3. Many efforts have aimed at addressing this deficiency. Among them, microfluidic technology has shown great advantages, by integrating RT-RPA and a universal LF detection system into a single chip. Results can be obtained with only simple nucleic acid extraction, loading, and incubation for approximately 30 minutes. This MI-IF-RPA detection method is rapid and sensitive, and it effectively decreases the risk of contamination [74].

TABLE 3 |

Comparison of RPA types.

RPA types	Advantages	Disadvantages
RPA-LF	1. Results can be obtained extremely rapidly with a visual read-out 2. Testing equipment is simple and suitable for resource-limited or non-laboratory environments	1. Insufficient accuracy and stability 2. Proneness to environmental contamination and false positives
Real-time fluorescence RPA	1. Shorter detection time 2. Closed reaction tubes decrease the risk of contamination during operation	Need for a thermostatic fluorescence detection instrument limits ease of use
CRISPR/Cas-RPA	1. The LOD is very low 2. The components of the CRISPR and RPA assays have similar reaction temperatures (37°C), and reactions can be performed in one tube	1. Long detection time 2. Additional labeled ssDNA or ssRNA reporter increases the cost
Electrochemical RPA	Solid phase reaction enables the integration of amplification, hybridization, and detection on a platform, thus decreasing analysis time and contamination	Lower amplification efficiency due to solid phase reaction leads

Real-time fluorescence RPA

As with PCR, RPA amplification results can be monitored by real-time fluorescence [75]. Fluorophore dyes, such as SYBR Green and Eva Green, can be used for real time detection [76,77]. However, these dyes cannot distinguish between amplicons and primer dimers, thus potentially leading to false positive results. Therefore, specific probes are preferred in RPA reactions, including the Exo and Fpg probes, named after the corresponding enzymes [75,78]. The Exo probe carries a fluorescence group and a fluorescence quenching group which are separated by a tetrahydrofuran (THF) residue [79]. During RPA amplification, the DNA repair enzyme exonuclease III cleaves the tetrahydrofuran in the Exo probe, thus leading to separation of the fluorescence group and the fluorescence quenching group, and facilitating the generation of fluorescence that is subsequently monitored.

Generally, real-time fluorescence RPA detection requires less time to complete than RPA-LF. In addition, the lids of the reaction tubes need not be kept open, thereby decreasing the risk of contamination during operation. The only disadvantage of this method is the requirement for thermostatic fluorescence detection instrument, which may limit its ease of use Table 3. The low reaction temperature is advantageous for miniaturization because of the low energy input required; therefore this method is a favorable candidate for battery powered hand-held devices [12]. Researchers have attempted to design and produce simple portable fluorescent readers [80,81].

Ultrasensitive real-time fluorescence RPA methods have been established for the detection of zoonotic pathogens. The LOD for detecting the kinetoplast minicircle DNA of Leishmania donovani [80], the CeuE gene of Campylobacter jejuni [82], the hipO gene of Campylobacter coli [82], and the 18S RNA gene of Plasmodium knowlesi [83] has been found to be as low as 1 cell/reaction, 1 CFU/ml, 1 CFU/ml, and 1 plasmid/reaction, respectively.

Euler et al. have developed ten detection methods for eight zoonotic pathogens that are also biothreat agents: Francisella tularensis, Yersinia pestis, Bacillus anthracis, and variola virus, by using RPA assays, and Rift Valley fever virus, Ebola virus, Sudan virus, and Marburg virus by using RT-RPA assays [12]. The analytical sensitivity ranges from 16 to 21 molecules detected, and the detection time ranges from 4 to 10 minutes—a detection performance surpassing those of PCR, real-time PCR, or LAMP [12].

Other zoonotic pathogens for which real-time fluorescence RPA detection has been successfully implemented include Streptococcus suis serotype 2, Mycobacterium tuberculosis, Rickettsia spp., yellow fever virus, dengue virus types 1–4, orf virus, rabies virus, avian influenza virus, hepatitis E virus, Chikungunya virus, Crimean-Congo hemorrhagic fever virus, Zika virus, highly pathogenic porcine reproductive and respiratory syndrome virus, and SARS-CoV-2 [30,48,81,84–94].

Now that various detection methods for single zoonotic pathogens have been established, integrating multiple methods to produce multiplex detection reagents or devices is a major development trend for fluorescence RPA. Researchers have attempted to integrate multiple methods into microfluidic chips to detect dozens of pathogens simultaneously [95,96]. In addition, a mobile suitcase laboratory for detecting zoonotic pathogens in the field and has been used in several studies [80,81,90,97].

CRISPR/Cas-RPA detection

One constraint of isothermal amplification is that single-nucleotide polymorphisms, which are crucial in both pathogen and disease detection, cannot always be discriminated [47,98]. A new molecular diagnostic tool based on the CRISPR/Cas system can overcome this weakness [33]. The origin of CRISPR based detection was rooted in the discovery of the collateral cleavage activity of protein Cas13a. When Cas13a binds CRISPR RNA (crRNA), the crRNA specifically pairs with a target sequence and subsequently induces both enzymatic cleavage of both the targeted sequence and untargeted collateral cleavage of all single stranded RNA (ssRNA) [99]. A comprehensive CRISPR system called SHERLOCK, which combines the CRISPR/Cas system with RPA [33], relies on the collateral trans-cleavage of quenched fluorescent nucleotides after target binding. Over the years, other Cas proteins, including Cas9, Cas12, and Cas14, have also been demonstrated to function in DNA or RNA sensing with high sensitivity and selectivity [47]. Myhrvold et al. have developed a complement to SHERLOCK called HUDSON technology to detect viruses directly from body fluids [100]. Chen et al. have used Cas12a collateral trans-cleavage and isothermal amplification to develop the DETECTR method [101], which can achieve amol/L sensitivity for DNA detection. Li et al. have developed a highly sensitive nucleic acid detection method called HOLMES by using Cas12a and an ssDNA fluorescence probe for rapid detection of DNA and RNA viruses with a sensitivity as low as 1–10 amol/L [102].

Currently, although signal amplification has been improved by introducing spherical nucleic acid reporters or multiple crRNAs [103,104], the sensitivity still does not meet the requirements for clinical detection, and the method is not suitable for application without a nucleotide amplification procedure. To amplify the signal and improve the detection sensitivity, CRISPR/Cas diagnostic technology is usually combined with NAA technology, such as PCR, LAMP, or RPA. Compared with other NAA assays, RPA has an inherent advantage in its compatibility with the CRISPR/Cas system, because both methods use reaction temperatures around 37°C. On the basis of this feature, the All-In-One Dual CRISPR-Cas12a (AIOD-CRISPR) assay for one-pot, ultrasensitive, and visual SARS-CoV-2 detection has been developed [105,106], in which, the components for both RPA and CRISPR-based detection are prepared in one pot, thus avoiding separate pre-amplification of target nucleic acids [101], or physical separation of the Cas enzyme [107].

In the detection of other zoonotic pathogens, to our knowledge, only limited studies have used CRISPR/Cas12a or CRISPR/Cas13a for bacteria or viruses, such as Leptospira, Salmonella spp., Zika virus, dengue virus, avian influenza A (H7N9) virus, influenza A virus, influenza B virus, and rabies virus [100,108–111].

The limited use of this technology in the detection of zoonotic pathogens may be due to its short duration of application. In addition, compared with RPA-LF and real-time fluorescence RPA, CRISPR/Cas-RPA detection uses additional labeled ssDNA or ssRNA reporter for collateral cleavage, thus potentially increasing the cost Table 3. The advantage of the CRISPR/Cas-RPA detection method in discriminating single-nucleotide polymorphisms may become a disadvantage when mutations are present in the target sequences in clinical applications, which focus on disease diagnosis rather than typing. Thus, this novel and promising detection method requires more studies to support applications in zoonotic pathogen detection.

Electrochemical RPA

Electrochemical RPA detection relies on the rapid isothermal amplification of target pathogen DNA sequences by RPA followed by gold nanoparticle-based electrochemical assessment with differential pulse voltammetry. This method couples RPA and electrochemistry on disposable screen printed carbon electrodes, and electrochemically active substances are used to generate signals associated with NAA [112].

In the detection of zoonotic pathogens, a rapid electrochemical detection method for Mycobacterium tuberculosis based on colloidal gold nanoparticles has been reported with a LOD as low as 1 CFU [113]. del Río et al. have constructed an electrochemical platform with a lower LOD of 1×10⁻¹⁵ M for Francisella tularensis detection [51]. In addition, the electrochemical RPA in detection of SARS-CoV-2 by using human body temperature was established and had a LOD slightly below or comparable to that of RPA assay results obtained from gel electrophoresis without post-amplification purification [114]. The method usually uses a solid phase and requires a handheld device for electrochemical measurement in on-site detection. The solid-phase RPA approach, on the one hand, allows for integration of DNA amplification, hybridization, and detection on a platform, thus decreasing analysis time and contamination, and potentially enabling on-site testing, but on the other hand, leads to lower amplification efficiency than that in solution, owing to steric hindrance effects on various components in the amplification system Table 3 [115]. However, the electrochemical RPA remains under development, and more work is required to improve its performance and capabilities [115].

Other methods coupled with RPA

Other detection methods coupled with RPA have been reported for zoonotic pathogen detection, including flocculation assays, chemiluminescence, and silicon microring resonator (SMR)-based photonic detection [16].

Flocculation analysis detection is based on the phenomenon of colloidal chemical bridging flocculation. The flocculation assay detection was first combined with an RPA assay reported by Wee et al., in which RPA amplicons on the magnetic bead surfaces cross-link multiple other RPA-magnetic bead conjugates, thus causing a sharp transition from solution phase to flocculate [116], which is detectable by the naked eye. The method was subsequently extended to detect zoonotic pathogens including malaria parasites, Mycobacterium tuberculosis, and the influenza virus H1N1 [116,117].

The chemiluminescent detection converts chemical energy into the emission of visible light, as the result of an oxidation or hydrolysis reaction [16]. RPA assays coupled with chemiluminescent detection for several zoonotic pathogens, including HAdV 41, Legionella spp. and Legionella pneumophila, have been applied on flow-based microarrays [118,119]. This method can be used for multiplex detection after immobilization of one of the two primers from different pathogens on one chip for asymmetric amplification. However, the procedure is tedious and time-consuming, thus limiting its use in field applications and resource-limited regions.

SMR-based photonic detection also involves performing NAA in an asymmetric manner. One primer is pre-immobilized on the SMR, and the binding of nucleic acids to the pre-immobilized primer induces changes in the refractive index proximal to the waveguide surface, which can be monitored in real time through the SMR. Applications in the detection of zoonotic pathogens including M. tuberculosis and F. tularensis, have demonstrated that SMR-RPA is an alternative detection method to fluorophore-based real-time detection, with the advantages of being label-free and much more sensitive [120,121].

Applications of RPA in SARS-CoV-2 detection

SARS-CoV-2 has become a serious public health concern in recent years [122,123]. Because many infected patients are asymptomatic, the total number of infections remains unclear [124]. The development of sensitive, rapid, specific, and cost-effective detection methods has never been more important. RPA has been widely used for the detection of SARS-CoV-2 viral RNA in clinical samples [71,97,125]. Combined with LF assays, RT-RPA can detect pathogenic nucleic acids within 20 minutes, with a detection limit as low as 7.659 copies/μL RNA [69]. With a microfluidic chip that integrates RT-RPA and universal LF, the detection limit can be increased to 1 copy per/μl, with an incubation time of approximately 30 minutes [74].

The CRISPR-Cas system has recently been used to sensitively detect nucleic acids, and numerous CRISPR-Cas-RPA detection systems have been developed [125–128]. Most of these methods are based on a combination of CRISPR/Cas12a and RT-RPA, with the introduction of a fluorescence probe for fluorescence readout or gold nanoparticles for colorimetric readout [31,122,126,127,129–132]. For example, a CRISPR–Cas12-based assay combined with DNA-modified gold nanoparticles has been developed, with a detection limit of one viral genome sequence copy per test. However, the detection time is increased to 50 minutes, 30 minutes of which are used for colorimetric readings [122,126,127]. Two separate teams have combined CRISPR/Cas9, LF assays, and RT-RPA technology as a platform for visual detection of SARS-CoV-2, thus providing an accurate and convenient pathway for diagnosis of COVID-19 or other infectious diseases in resource-limited regions [128,133]. Few studies have used CRISPR/Cas13a, in which an additional transcription step is needed and may increase the detection time. Arizti-Sanz et al. have identified the optimal conditions to allow single-step Cas13-based detection and RPA, and have developed a sensitive and specific diagnostic tool that can detect SARS-CoV-2 RNA from unextracted samples, with a sample-to-answer time of 50?minutes [134]. Moreover, Tian et al. have designed a system using both Cas12a and Cas13a for dual-gene detection, in which, dual-gene amplified products from the multiplex RPA are simultaneously detected by Cas12a and Cas13a assays in a single tube [135].

In addition to LF assays and the CRISPR-Cas system, many other detection techniques have been combined with RPAs. Wang et al. have established a ligation and recombinase polymerase amplification method. Using a high concentration of T4 DNA ligase, this method has achieved a satisfactory sensitivity of ten copies per reaction within 30 minutes [136]. Furthermore, Moon et al. have combined an rkDNA-graphene oxide probe system with RPA and developed a rapid detection method with extremely high sensitivity (LOD 6.0 aM) [137].

At present, with the continual development of the COVID-19 pandemic, research on rapid and sensitive detection of the SARS-CoV-2 virus remains a development focus.

CONCLUSIONS AND FUTURE PERSPECTIVES

In recent years, zoonoses have caused great economic losses worldwide and severely threatened human health, life, and safety. Because traditional detection technologies can no longer meet the detection requirements for zoonotic pathogens, rapid, sensitive, specific, multiplex detection methods must be established. As an emerging molecular detection technology, RPA assays have been widely used in medicine and pharmacy applications, and also have begun to emerge in the detection of zoonotic pathogens. RPA assays have many technical advantages: they do not require thermal cycling, and the reaction can be completed at relatively lower temperatures of 37–42 °C; the reaction is fast, with amplification times of 5–20 minutes; the method is portable, and combinations of RPA and LF, fluorescence, the CRISPR/Cas system, and other technologies have been achieved. Since the COVID-19 pandemic, RPA technology has been crucial for rapid pathogen detection.

However, RPA assays are novel, and their use has not been as widespread or common as PCR methods in the detection of zoonotic pathogens, although these methods have developed faster since the COVID-19 pandemic. Because RPA has the advantage of being naturally suitable for on-site testing, more attention is needed in integrating sample preparation with RPA detection to achieve a fast sample-to-result pipeline that would enable a complete RPA assay for on-site or field application. Multiplex high-throughput detection is another research and application direction for zoonotic pathogen detection. In this respect, the combination of microfluid or microarray technology with RPA assay shows good prospects. In addition, as suggested by Li et al. [16], developing wearable sensors and performing RPA assays using human body heat to detect potential zoonotic pathogens may revolutionize RPA diagnostics to enable self-testing. With its continual rapid development, RPA is expected to play a more important role in the prevention and control of zoonotic diseases in the near future, particularly in mobile and point-of-care applications.

CONFLICT OF INTEREST

The authors declare that they have no competing interests.

REFERENCES

1. Saiki RK, Bugawan TL, Horn GT, Mullis KB, Erlich HA. Analysis of enzymatically amplified beta-globin and HLA-DQ alpha DNA with allele-specific oligonucleotide probes. Nature. 1986. Vol. 324(6093):163–166

2. Craw P, Balachandran W. Isothermal nucleic acid amplification technologies for point-of-care diagnostics: a critical review. Lab Chip. 2012. Vol. 12(14):2469–2486

3. Deng H, Gao Z. Bioanalytical applications of isothermal nucleic acid amplification techniques. Anal Chim Acta. 2015. Vol. 853:30–45

4. Lei Y, Jie Z, Yue Z, Garrison AS, Roembke BT, Nakayama S, et al.. Isothermal amplified detection of DNA and RNA. Mol Biosyst. 2014. Vol. 10(5):970–1003

5. Li J, Macdonald J. Advances in isothermal amplification: novel strategies inspired by biological processes. Biosens Bioelectron. 2015. Vol. 64:196–211

6. Zhao Y, Chen F, Li Q, Wang L, Fan C. Isothermal amplification of nucleic acids. Chem Rev. 2015. Vol. 115(22):12491–12545

7. Piepenburg O, Williams CH, Stemple DL, Armes NA. DNA detection using recombination proteins. PLoS Biol. 2006. Vol. 4(7):1115–1121

8. Euler M, Wang Y, Otto P, Tomaso H, Escudero R, Anda P, et al.. Recombinase polymerase amplification assay for rapid detection of Francisella tularensis . J Clin Microbiol. 2012. Vol. 50(7):2234–2238

9. Kojima K, Juma KM, Akagi S, Hayashi K, Takita T, O’Sullivan CK, et al.. Solvent engineering studies on recombinase polymerase amplification. J Biosci Bioeng. 2020. Vol. 131(2):219–224

10. Mcquillan JS, Wilson MW. Recombinase polymerase amplification for fast, selective, DNA-based detection of faecal indicator Escherichia coli . Lett Appl Microbiol. 2021. Vol. 72(4):382–389

11. Wee EJ, Ngo TH, Trau M. Colorimetric detection of both total genomic and loci-specific DNA methylation from limited DNA inputs. Clin Epigenet. 2015. Vol. 7(1):65

12. Euler M, Wang Y, Heidenreich D, Patel P, Strohmeier O, Hakenberg S, et al.. Development of a panel of recombinase polymerase amplification assays for detection of biothreat agents. J Clin Microbiol. 2013. Vol. 51(4):1110–1117

13. Mekuria TA, Zhang S, Eastwell KC. Rapid and sensitive detection of Little cherry virus 2 using isothermal reverse transcription-recombinase polymerase amplification. J Virol Methods. 2014. Vol. 205:24–30

14. Wee EJH, Trau M. Simple isothermal strategy for multiplexed, rapid, sensitive, and accurate miRNA detection. ACS Sens. 2016. Vol. 1(6):670–675

15. Wang J, Wang J, Geng Y, Yuan W. A recombinase polymerase amplification-based assay for rapid detection of African swine fever virus. Can J Vet Res. 2017. Vol. 81(4):308–312

16. Li J, Macdonald J, Stetten FV. Review: A comprehensive summary of a decade development of the recombinase polymerase amplification. Analyst. 2018. Vol. 144(1):31–67

17. Euler M, Wang Y, Nentwich O, Piepenburg O, Hufert FT, Weidmann M. Recombinase polymerase amplification assay for rapid detection of Rift Valley fever virus. J Clin Virol. 2012. Vol. 54(4):308–312

18. Jarvi SI, Atkinson ES, Kaluna LM, Snook KA, Steel A. Development of a recombinase polymerase amplification (RPA-EXO) and lateral flow assay (RPA-LFA) based on the ITS1 gene for the detection of Angiostrongylus cantonensis in gastropod intermediate hosts. Parasitology. 2021. Vol. 148(2):251–258

19. Qi Y, Yin Q, Shao Y, Cao M, Li S, Chen H, et al.. Development of a rapid and visual nucleotide detection method for a Chinese epidemic strain of Orientia tsutsugamushi based on recombinase polymerase amplification assay and lateral flow test. Int J Infect Dis. 2018. Vol. 70:42–50

20. Qi Y, Li W, Li X, Shen W, Zhang J, Li J, et al.. Development of rapid and visual nucleic acid detection methods towards four serotypes of human adenovirus species B based on RPA-LF test. Biomed Res Int. 2021. Vol. 2021:9957747

21. Qi Y, Shao Y, Rao J, Shen W, Yin Q, Li X, et al.. Development of a rapid and visual detection method for Rickettsia rickettsii combining recombinase polymerase assay with lateral flow test. PLoS One. 2018. Vol. 13(11):e0207811

22. Qi Y, Yin Q, Shao Y, Li S, Chen H, Shen W, et al.. Rapid and visual detection of Coxiella burnetii using recombinase polymerase amplification combined with lateral flow strips. Biomed Res Int. 2018. Vol. 2018:6417354

23. Kersting S, Rausch V, Bier FF, Nickisch-Rosenegk MV. Rapid detection of Plasmodium falciparum with isothermal recombinase polymerase amplification and lateral flow analysis. Malar J. 2014. Vol. 13(1):1–9

24. Chandu D, Paul S, Parker M, Dudin Y, King-Sitzes J, Perez T, et al.. Development of a rapid point-of-use DNA test for the screening of genuity roundup ready 2 yield soybean in seed samples. Biomed Res Int. 2016. Vol. 2016:3145921

25. Fuller SL, Savory EA, Weisberg AJ, Buser JZ, Gordon MI, Putnam M, et al.. Isothermal amplification and lateral-flow assay for detecting crown-gall-causing Agrobacterium spp. Phytopathology. 2017. Vol. 107(9):1062–1068

26. Lillis L, Lehman D, Singhal MC, Cantera J, Singleton J, Labarre P, et al.. Non-instrumented incubation of a recombinase polymerase amplification assay for the rapid and sensitive detection of proviral HIV-1 DNA. PLoS One. 2014. Vol. 9(9):e108189

27. Xia X, Yu Y, Weidmann M, Pan Y, Yan S, Wang Y. Rapid detection of shrimp white spot syndrome virus by real time, isothermal recombinase polymerase amplification assay. PLoS One. 2014. Vol. 9(8):e104667

28. Chao CC, Belinskaya T, Zhang Z, Ching WM. Development of recombinase polymerase amplification assays for detection of Orientia tsutsugamushi or Rickettsia typhi . PLoS Negl Trop Dis. 2015. Vol. 9(7):e0003884

29. Wang J, Wang J, Li R, Liu L, Yuan W. Rapid and sensitive detection of canine distemper virus by real-time reverse transcription recombinase polymerase amplification. BMC Vet Res. 2017. Vol. 13(1):241

30. Behrmann O, Bachmann I, Spiegel M, Schramm M, Abd El Wahed A, Dobler G, et al.. Rapid detection of SARS-CoV-2 by low volume real-time single tube reverse transcription recombinase polymerase amplification using an exo probe with an Internally Linked Quencher (Exo-IQ). Clin Chem. 2020. Vol. 66(8):1047–1054

31. Feng W, Peng H, Xu J, Liu Y, Pabbaraju K, Tipples G, et al.. Integrating reverse transcription recombinase polymerase amplification with CRISPR technology for the one-tube assay of RNA. Anal Chem. 2021. Vol. 93(37):12808–12816

32. Lalremruata A, Nguyen TT, McCall MBB, Mombo-Ngoma G, Agnandji ST, Adegnika AA, et al.. Recombinase polymerase amplification and lateral flow assay for ultrasensitive detection of low-density plasmodium falciparum infection from controlled human malaria infection studies and naturally acquired infections. J Clin Microbiol. 2020. Vol. 58(5):e01879–19

33. Gootenberg JS, Abudayyeh OO, Lee JW, Essletzbichler P, Dy AJ, Joung J, et al.. Nucleic acid detection with CRISPR-Cas13a/C2c2. Science. 2017. Vol. 356(6336):438–442

34. Mccoy AG, Miles TD, Bilodeau GJ, Woods P, Blomquist C, Martin FN, et al.. Validation of a preformulated, field deployable, recombinase polymerase amplification assay for Phytophthora species. Plants. 2020. Vol. 9(4):466

35. Rohrman BA, Richards-Kortum RR. A paper and plastic device for performing recombinase polymerase amplification of HIV DNA. Lab Chip. 2012. Vol. 12(17):3082–3088

36. Ma B, Li J, Chen K, Yu X, Sun C, Zhang M. Multiplex recombinase polymerase amplification assay for the simultaneous detection of three foodborne pathogens in seafood. Foods. 2020. Vol. 9(3):278

37. Frimpong M, Simpson SV, Ahor HS, Agbanyo A, Gyabaah S, Agbavor B, et al.. Multiplex recombinase polymerase amplification assay for simultaneous detection of Treponema pallidum and Haemophilus ducreyi in yaws-like lesions. Trop Med Infect Dis. 2020. Vol. 5(4):157

38. Tsai SK, Chen CC, Lin HJ, Lin HY, Chen TT, Wang LC. Combination of multiplex reverse transcription recombinase polymerase amplification assay and capillary electrophoresis provides high sensitive and high-throughput simultaneous detection of avian influenza virus subtypes. J Vet Sci. 2020. Vol. 21(2):e24

39. Cross AR, Baldwin VM, Roy S, Essex-Lopresti AE, Prior JL, Harmer NJ. Zoonoses under our noses. Microbes Infect. 2019. Vol. 21(1):10–19

40. Taylor LH, Latham SM, Woolhouse ME. Risk factors for human disease emergence. Philos Trans R Soc Lond B Biol Sci. 2001. Vol. 356(1411):983–989

41. Fasanella A, Galante D, Garofolo G, Jones MH. Anthrax undervalued zoonosis. Vet Microbiol. 2010. Vol. 140(3-4):318–331

42. Mortimer PP. Influenza: the centennial of a zoonosis. Rev Med Virol. 2019. Vol. 29(1):e2030

43. Wilde H, Hemachudha T, Wacharapluesadee S, Lumlertdacha B, Tepsumethanon V. Rabies in Asia: the classical zoonosis. Curr Topics Microbiol Immunol. 2013. Vol. 365:185–203

44. Gebreyes WA, Dupouy-Camet J, Newport MJ, Oliveira CJ, Schlesinger LS, Saif YM, et al.. The global one health paradigm: challenges and opportunities for tackling infectious diseases at the human, animal, and environment interface in low-resource settings. PLoS Negl Trop Dis. 2014. Vol. 8(11):e3257

45. Cascio A, Bosilkovski M, Rodriguez-Morales AJ, Pappas G. The socio-ecology of zoonotic infections. Clin Microbiol Infect. 2011. Vol. 17(3):336–342

46. Bennett R, Ijpelaar J. Updated estimates of the costs associated with thirty four endemic livestock diseases in great Britain: a note. J Agric Econ. 2010. Vol. 56(1):135–144

47. van Dongen JE, Berendsen JTW, Steenbergen RDM, Wolthuis RMF, Eijkel JCT, Segerink LI. Point-of-care CRISPR/Cas nucleic acid detection: recent advances, challenges and opportunities. Biosens Bioelectron. 2020. Vol. 166:112445

48. Xu Y, Wu P, Zhang H, Li J. Rapid detection of Mycobacterium tuberculosis based on antigen 85B via real-time recombinase polymerase amplification. Lett Appl Microbiol. 2021. Vol. 72(2):106–112

49. Gumaa MM, Cao X, Li Z, Lou Z, Zhang N, Zhang Z, et al.. Establishment of a recombinase polymerase amplification (RPA) assay for the detection of Brucella spp . Infection. Mol Cell Probes. 2019. Vol. 47:101434

50. Chen G, Lyu Y, Wang D, Zhu L, Cao S, Pan C, et al.. Obtaining specific sequence tags for Yersinia pestis and visually detecting them using the CRISPR-Cas12a system. Pathogens. 2021. Vol. 10(5):562

51. Del Rio JS, Lobato IM, Mayboroda O, Katakis I, O’Sullivan CK. Enhanced solid-phase recombinase polymerase amplification and electrochemical detection. Anal Bioanal Chem. 2017. Vol. 409(12):3261–3269

52. Liu W, Liu HX, Zhang L, Hou XX, Wan KL, Hao Q. A novel isothermal assay of Borrelia burgdorferi by recombinase polymerase amplification with lateral flow detection. Int J Mol Sci. 2016. Vol. 17(8):1250

53. Rani A, Ravindran VB, Surapaneni A, Shahsavari E, Haleyur N, Mantri N, et al.. Evaluation and comparison of recombinase polymerase amplification coupled with lateral-flow bioassay for Escherichia coli O157:H7 detection using diifeerent genes. Sci Rep. 2021. Vol. 11(1):1881

54. Faye M, Abd El Wahed A, Faye O, Kissenkotter J, Hoffmann B, Sall AA, et al.. A recombinase polymerase amplification assay for rapid detection of rabies virus. Sci Rep. 2021. Vol. 11(1):3131

55. Ma S, Li X, Peng B, Wu W, Wang X, Liu H, et al.. Rapid detection of avian influenza A virus (H7N9) by lateral flow dipstick recombinase polymerase amplification. Biol Pharm Bull. 2018. Vol. 41(12):1804–1808

56. Yang M, Ke Y, Wang X, Ren H, Liu W, Lu H, et al.. Development and evaluation of a rapid and sensitive EBOV-RPA test for rapid diagnosis of Ebola virus disease. Sci Rep. 2016. Vol. 6:26943

57. Xi Y, Xu CZ, Xie ZZ, Zhu DL, Dong JM. Rapid and visual detection of dengue virus using recombinase polymerase amplification method combined with lateral flow dipstick. Mol Cell Probes. 2019. Vol. 46:101413

58. Vasileva Wand NI, Bonney LC, Watson RJ, Graham V, Hewson R. Point-of-care diagnostic assay for the detection of Zika virus using the recombinase polymerase amplification method. J Gen Virol. 2018. Vol. 99(8):1012–1026

59. Posthuma-Trumpie GA, Korf J, Amerongen AV. Lateral flow (immuno)assay: its strengths, weaknesses, opportunities and threats. a literature survey. Anal Bioanal Chem. 2009. Vol. 393(2):569–582

60. Cordray MS, Richards-Kortum RR. A paper and plastic device for the combined isothermal amplification and lateral flow detection of Plasmodium DNA. Malar J. 2015. Vol. 14:472

61. Salazar A, Ochoa-Corona FM, Talley JL, Noden BH. Recombinase polymerase amplification (RPA) with lateral flow detection for three Anaplasma species of importance to livestock health. Sci Rep. 2021. Vol. 11(1):15962

62. Wu YD, Xu MJ, Wang QQ, Zhou CX, Wang M, Zhu XQ, et al.. Recombinase polymerase amplification (RPA) combined with lateral flow (LF) strip for detection of Toxoplasma gondii in the environment. Vet Parasitol. 2017. Vol. 243:199–203

63. Jimenez-Coello M, Shelite T, Castellanos-Gonzalez A, Saldarriaga O, Rivero R, Ortega-Pacheco A, et al.. Efficacy of recombinase polymerase amplification to diagnose Trypanosoma cruzi infection in dogs with cardiac alterations from an endemic area of Mexico. Vector Borne Zoonotic Dis. 2018. Vol. 18(8):417–423

64. Saxena A, Pal V, Tripathi NK, Goel AK. Development of a rapid and sensitive recombinase polymerase amplification-lateral flow assay for detection of Burkholderia mallei . Transbound Emerg Dis. 2019. Vol. 66(2):1016–1022

65. Castellanos-Gonzalez A, Saldarriaga OA, Tartaglino L, Gacek R, Temple E, Sparks H, et al.. A novel molecular test to diagnose canine visceral leishmaniasis at the point of care. Am J Trop Med Hyg. 2015. Vol. 93(5):970–975

66. El-Tholoth M, Branavan M, Naveenathayalan A, Balachandran W. Recombinase polymerase amplification-nucleic acid lateral flow immunoassays for Newcastle disease virus and infectious bronchitis virus detection. Mol Biol Rep. 2019. Vol. 46(6):6391–6397

67. Krõlov K, Frolova J, Tudoran O, Suhorutsenko J, Lehto T, Sibul H, et al.. Sensitive and rapid detection of Chlamydia trachomatis by recombinase polymerase amplification directly from urine samples. J Mol Diagn. 2014. Vol. 16(1):127–135

68. Yang Y, Qin X, Wang G, Jin J, Shang Y, Zhang Z. Development of an isothermoal amplification-based assay for rapid visual detection of an Orf virus. Virol J. 2016. Vol. 13:46

69. Lau YL, Ismail IB, Mustapa NIB, Lai MY, Tuan Soh TS, Haji Hassan A, et al.. Development of a reverse transcription recombinase polymerase amplification assay for rapid and direct visual detection of Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2). PLoS One. 2021. Vol. 16(1):e0245164

70. Wang Z, Yang PP, Zhang YH, Tian KY, Bian CZ, Zhao J. Development of a reverse transcription recombinase polymerase amplification combined with lateral-flow dipstick assay for avian influenza H9N2 HA gene detection. Transbound Emerg Dis. 2019. Vol. 66(1):546–551

71. Shelite TR, Uscanga-Palomeque AC, Castellanos-Gonzalez A, Melby PC, Travi BL. Isothermal recombinase polymerase amplification-lateral flow detection of SARS-CoV-2, the etiological agent of COVID-19. J Virol Methods. 2021. Vol. 296:114227

72. Shelite TR, Bopp NE, Moncayo A, Reynolds ES, Thangamani S, Melby PC, et al.. Isothermal recombinase polymerase amplification-lateral flow point-of-care diagnostic test for Heartland virus. Vector Borne Zoonotic Dis. 2021. Vol. 21(2):110–115

73. Kasetsirikul S, Shiddiky M, Nguyen NT. Challenges and perspectives in the development of paper-based lateral flow assays. Microfluid Nanofluid. 2020. Vol. 24(2):17.11–17.18

74. Liu D, Shen H, Zhang Y, Shen D, Zhu M, Song Y, et al.. A microfluidic-integrated lateral flow recombinase polymerase amplification (MI-IF-RPA) assay for rapid COVID-19 detection. Lab Chip. 2021. Vol. 21(10):2019–2026

75. Davi SD, Kissenkotter J, Faye M, Bohlken-Fascher S, Stahl-Hennig C, Faye O, et al.. Recombinase polymerase amplification assay for rapid detection of Monkeypox virus. Diagn Microbiol Infect Dis. 2019. Vol. 95(1):41–45

76. Lai MY, Lau YL. Detection of Plasmodium knowlesi using recombinase polymerase amplification (RPA) combined with SYBR Green I. Acta Tropica. 2020. Vol. 208:105511

77. Zhang S, Sun A, Wan B, Du Y, Wu Y, Zhang A, et al.. Development of a directly visualized recombinase polymerase amplification-SYBR green I method for the rapid detection of African swine fever virus. Front Microbiol. 2020. Vol. 11:602709

78. Pang Y, Cong F, Zhang X, Li H, Chang YF, Xie Q, et al.. A recombinase polymerase amplification-based assay for rapid detection of Chlamydia psittaci . Poult Sci. 2021. Vol. 100(2):585–591

79. Wang JC, Liu LB, Han QA, Wang JF, Yuan WZ. An exo probe-based recombinase polymerase amplification assay for the rapid detection of porcine parvovirus. J Virol Methods. 2017. Vol. 248:145–147

80. Mondal D, Ghosh P, Khan MAA, Hossain F, Böhlken-Fascher S, Matlashewski G, et al.. Mobile suitcase laboratory for rapid detection of Leishmania donovani using recombinase polymerase amplification assay. Parasit Vectors. 2016. Vol. 9(1):281

81. Kissenkotter J, Hansen S, Bohlken-Fascher S, Ademowo OG, Oyinloye OE, Bakarey AS, et al.. Development of a pan-rickettsial molecular diagnostic test based on recombinase polymerase amplification assay. Anal Biochem. 2018. Vol. 544:29–33

82. Kim JY, Lee JL. Development of a multiplex real-time recombinase polymerase amplification (RPA) assay for rapid quantitative detection of Campylobacter coli and jejuni from eggs and chicken products. Food Control. 2016. Vol. 73:1247–1255

83. Lai MY, Ooi CH, Lau YL. Rapid detection of plasmodium knowlesi by isothermal recombinase polymerase amplification assay. Am J Trop Med Hyg. 2017. Vol. 97(5):1597–1599

84. Gao S, Wang J, Li D, Li Y, Lou C, Zha E, et al.. Development and evaluation of a time-saving RT-qRPA method for the detection of genotype 4 HEV presence in raw pork liver. Int J Food Microbiol. 2020. Vol. 322:108587

85. Coertse J, Weyer J, Nel LH, Markotter W. Reverse transcription recombinase polymerase amplification assay for rapid detection of canine associated rabies virus in Africa. PLoS One. 2019. Vol. 14(7):e0219292

86. Wang S, Huang B, Ma X, Liu P, Wang Y, Zhang X, et al.. Reverse-transcription recombinase-aided amplification assay for H7 subtype avian influenza virus. Transbound Emerg Dis. 2020. Vol. 67(2):877–883

87. Jiang X, Zhu L, Zhan D. Development of a recombinase polymerase amplification assay for rapid detection of Streptococcus suis type 2 in nasopharyngeal swab samples. Diagn Microbiol Infect Dis. 2021. Vol. 102(2):115594

88. Patel P, Abd El Wahed A, Faye O, Prüger P, Kaiser M, Thaloengsok S, et al.. A field-deployable reverse transcription recombinase polymerase amplification assay for rapid detection of the Chikungunya Virus. PLoS Negl Trop Dis. 2016. Vol. 10(9):e0004953

89. Escadafal C, Faye O, Sall AA, Faye O, Weidmann M, Strohmeier O, et al.. Rapid molecular assays for the detection of yellow fever virus in low-resource settings. PLoS Negl Trop Dis. 2014. Vol. 8(3):e2730

90. Abd El Wahed A, Patel P, Faye O, Thaloengsok S, Heidenreich D, Matangkasombut P, et al.. Recombinase polymerase amplification assay for rapid diagnostics of Dengue Infection. PLoS One. 2015. Vol. 10(6):e0129682

91. Yang Y, Qin X, Wang G, Zhang Y, Shang Y, Zhang Z. Development of a fluorescent probe-based recombinase polymerase amplification assay for rapid detection of Orf virus. Virol J. 2015. Vol. 12:206

92. Yang Y, Qin X, Sun Y, Chen T, Zhang Z. Rapid detection of highly pathogenic porcine reproductive and respiratory syndrome virus by a fluorescent probe-based isothermal recombinase polymerase amplification assay. Virus Genes. 2016. Vol. 52(6):883–886

93. Bonney LC, Watson RJ, Afrough B, Mullojonova M, Dzhuraeva V, Tishkova F, et al.. A recombinase polymerase amplification assay for rapid detection of Crimean-Congo Haemorrhagic fever Virus infection. PLoS Negl Trop Dis. 2017. Vol. 11(10):e0006013

94. Tomar PS, Kumar S, Patel S, Kumar JS. Development and evaluation of real-time reverse transcription recombinase polymerase amplification assay for rapid and sensitive detection of West Nile virus in human clinical samples. Front Cell Infect Microbiol. 2021. Vol. 10:619071

95. Lutz S, Weber P, Focke M, Faltin B, Hoffmann J, Müller C, et al.. Microfluidic lab-on-a-foil for nucleic acid analysis based on isothermal recombinase polymerase amplification (RPA). Lab Chip. 2010. Vol. 10(7):887–893

96. Renner LD, Zan J, Hu LI, Martinez M, Resto PJ, Siegel AC, et al.. Detection of ESKAPE bacterial pathogens at the point of care using isothermal DNA-Based assays in a portable degas-actuated microfluidic diagnostic assay platform. Appl Environ Microbiol. 2017. Vol. 83(4):e02449–16

97. El Wahed AA, Patel P, Maier M, Pietsch C, Ruster D, Bohlken-Fascher S, et al.. Suitcase lab for rapid detection of SARS-CoV-2 based on recombinase polymerase amplification assay. Anal Chem. 2021. Vol. 93(4):2627–2634

98. Zanoli LM, Spoto G. Isothermal amplification methods for the detection of nucleic acids in microfluidic devices. Biosensors (Basel). 2013. Vol. 3(1):18–43

99. Abudayyeh OO, Gootenberg JS, Konermann S, Joung J, Slaymaker IM, Cox DBT, et al.. C2c2 is a single-component programmable RNA-guided RNA-targeting CRISPR effector. Science. 2016. Vol. 353(6299):aaf5573

100. Myhrvold C, Freije CA, Gootenberg JS, Abudayyeh OO, Metsky HC, Durbin AF, et al.. Field-deployable viral diagnostics using CRISPR-Cas13. Science. 2018. Vol. 360(6387):444–448

101. Chen JS, Ma E, Harrington LB, Da Costa M, Tian X, Palefsky JM, et al.. CRISPR-Cas12a target binding unleashes indiscriminate single-stranded DNase activity. Science. 2018. Vol. 360(6387):436–439

102. Li SY, Cheng QX, Wang JM, Li XY, Zhang ZL, Gao S, et al.. CRISPR-Cas12a-assisted nucleic acid detection. Cell Discov. 2018. Vol. 4:20

103. Fu X, Shi Y, Peng F, Zhou M, Yin Y, Tan Y, et al.. Exploring the trans-cleavage activity of CRISPR/Cas12a on gold nanoparticles for stable and sensitive biosensing. Anal Chem. 2021. Vol. 93(11):4967–4974

104. Fozouni P, Son S, Díaz de León Derby M, Knott GJ, Gray CN, D’Ambrosio MV, et al.. Amplification-free detection of SARS-CoV-2 with CRISPR-Cas13a and mobile phone microscopy. Cell. 2021. Vol. 184(2):323–333.e329

105. Ding X, Yin K, Li Z, Lalla RV, Ballesteros E, Sfeir MM, et al.. Ultrasensitive and visual detection of SARS-CoV-2 using all-in-one dual CRISPR-Cas12a assay. Nat Commun. 2020. Vol. 11(1):4711

106. Chen Y, Shi Y, Chen Y, Yang Z, Wu H, Zhou Z, et al.. Contamination-free visual detection of SARS-CoV-2 with CRISPR/Cas12a: a promising method in the point-of-care detection. Biosens Bioelectron. 2020. Vol. 169:112642

107. Wang B, Wang R, Wang D, Wu J, Li J, Wang J, et al.. Cas12aVDet: a CRISPR/Cas12a-based platform for rapid and visual nucleic acid detection. Anal Chem. 2019. Vol. 91(19):12156–12161

108. Jirawannaporn S, Limothai U, Tachaboon S, Dinhuzen J, Kiatamornrak P, Chaisuriyong W, et al.. Rapid and sensitive point-of-care detection of Leptospira by RPA-CRISPR/Cas12a targeting lipL32. PLoS Negl Trop Dis. 2022. Vol. 16(1):e0010112

109. An B, Zhang H, Su X, Guo Y, Wu T, Ge Y, et al.. Rapid and sensitive detection of Salmonella spp. using CRISPR-Cas13a combined with recombinase polymerase amplification. Front Microbiol. 2021. Vol. 12:732426

110. Liu Y, Xu H, Liu C, Peng L, Khan H, Cui L, et al.. CRISPR-Cas13a nanomachine based simple technology for avian influenza A (H7N9) virus on-site detection. J Biomed Nanotechnol. 2019. Vol. 15(4):790–798

111. Ren M, Mei H, Zhou J, Zhou M, Han H, Zhao L. Early diagnosis of rabies virus infection by RPA-CRISPR techniques in a rat model. Arch Virol. 2021. Vol. 166(4):1083–1092

112. Lau HY, Wu H, Wee EJH, Trau M, Wang Y, Botella JR. Specific and sensitive isothermal electrochemical biosensor for plant pathogen DNA detection with colloidal gold nanoparticles as probes. Sci Rep. 2017. Vol. 7:38896

113. Ng BYC, Xiao W, West NP, Wee EJ, Wang Y, Trau M. Rapid, single-cell electrochemical detection of Mycobacterium tuberculosis using colloidal gold nanoparticles. Anal Chem. 2015. Vol. 87(20):10613–10618

114. Kim HE, Schuck A, Lee SH, Lee Y, Kang M, Kim YS. Sensitive electrochemical biosensor combined with isothermal amplification for point-of-care COVID-19 tests. Biosens Bioelectron. 2021. Vol. 182:113168

115. Leonardo S, Toldrà A, Campàs M. Biosensors based on isothermal DNA amplification for bacterial detection in food safety and environmental monitoring. Sensors (Basel). 2021. Vol. 21(2):602

116. Wee EJH, Lau HY, Botella JR, Trau M. Re-purposing bridging flocculation for on-site, rapid, qualitative DNA detection in resource-poor settings. Chem Commun. 2015. Vol. 51(27):5828–5831

117. Ng BYC, Wee EJH, West NP, Trau M. Rapid DNA detection of mycobacterium tuberculosis-towards single cell sensitivity in point-of-care diagnosis. Sci Rep. 2015. Vol. 5:15027

118. Kober C, Niessner R, Seidel M. Quantification of viable and non-viable Legionella spp. by heterogeneous asymmetric recombinase polymerase amplification (haRPA) on a flow-based chemiluminescence microarray. Biosens Bioelectron. 2018. Vol. 100:49–55

119. Kunze A, Dilcher M, Abd El Wahed A, Hufert F, Niessner R, Seidel M. On-chip isothermal nucleic acid amplification on flow-based chemiluminescence microarray analysis platform for the detection of viruses and bacteria. Anal Chem. 2016. Vol. 88(1):898–905

120. Shin Y, Perera AP, Tang WY, Fu DL, Liu Q, Sheng JK, et al.. A rapid amplification/detection assay for analysis of Mycobacterium tuberculosis using an isothermal and silicon bio-photonic sensor complex. Biosens Bioelectron. 2015. Vol. 68:390–396

121. Sabaté Del Río J, Steylaerts T, Henry OYF, Bienstman P, Stakenborg T, Van Roy W, et al.. Real-time and label-free ring-resonator monitoring of solid-phase recombinase polymerase amplification. Biosens Bioelectron. 2015. Vol. 73:130–137

122. Xiong D, Dai W, Gong J, Li G, Liu N, Wu W, et al.. Rapid detection of SARS-CoV-2 with CRISPR-Cas12a. PLoS Biol. 2020. Vol. 18(12):e3000978

123. Wu F, Zhao S, Yu B, Chen YM, Wang W, Song ZG, et al.. A new coronavirus associated with human respiratory disease in China. Nature. 2020. Vol. 579(7798):265–269

124. Han D, Li R, Han Y, Zhang R, Li J. COVID-19: insight into the asymptomatic SARS-COV-2 infection and transmission. Int J Biol Sci. 2020. Vol. 16(15):2803–2811

125. Behrmann O, Bachmann I, Spiegel M, Schramm M, Abd El Wahed A, Dobler G, et al.. Rapid detection of SARS-CoV-2 by low volume real-time single tube reverse transcription recombinase polymerase amplification using an exo probe with an internally linked quencher (Exo-IQ). Clin Chem. 2020. Vol. 66(8):1047–1054

126. Huang Z, Tian D, Liu Y, Lin Z, Lyon CJ, Lai W, et al.. Ultra-sensitive and high-throughput CRISPR-p owered COVID-19 diagnosis. Biosens Bioelectron. 2020. Vol. 164:112316

127. Zhang WS, Pan J, Li F, Zhu M, Xu M, Zhu H, et al.. Reverse transcription recombinase polymerase amplification coupled with CRISPR-Cas12a for facile and highly sensitive colorimetric SARS-CoV-2 detection. Anal Chem. 2021. Vol. 93(8):4126–4133

128. Xiong E, Jiang L, Tian T, Hu M, Yue H, Huang M, et al.. Simultaneous dual-gene diagnosis of SARS-CoV-2 based on CRISPR/Cas9-mediated lateral flow assay. Angew Chem Int Ed Engl. 2021. Vol. 60(10):5307–5315

129. Mayuramart O, Nimsamer P, Rattanaburi S, Chantaravisoot N, Khongnomnan K, Chansaenroj J, et al.. Detection of severe acute respiratory syndrome coronavirus 2 and influenza viruses based on CRISPR-Cas12a. Exp Biol Med (Maywood). 2021. Vol. 246(4):400–405

130. Sun Y, Yu L, Liu C, Ye S, Chen W, Li D, et al.. W. One-tube SARS-CoV-2 detection platform based on RT-RPA and CRISPR/Cas12a. J Transl Med. 2021. Vol. 19(1):74

131. Yin K, Ding X, Li Z, Sfeir MM, Ballesteros E, Liu C. Autonomous lab-on-paper for multiplexed, CRISPR-based diagnostics of SARS-CoV-2. Lab Chip. 2021. Vol. 21(14):2730–2737

132. Talwar CS, Park KH, Ahn WC, Kim YS, Kwon OS, Yong D, et al.. Detection of infectious viruses using CRISPR-Cas12-based assay. Biosensors (Basel). 2021. Vol. 11(9):301

133. Marsic T, Ali Z, Tehseen M, Mahas A, Hamdan S, Mahfouz M. Vigilant: an engineered VirD2-Cas9 complex for lateral flow assay-based detection of SARS-CoV2. Nano Lett. 2021. Vol. 21(8):3596–3603

134. Arizti-Sanz J, Freije CA, Stanton AC, Petros BA, Boehm CK, Siddiqui S, et al.. Streamlined inactivation, amplification, and Cas13-based detection of SARS-CoV-2. Nat Commun. 2020. Vol. 11(1):5921

135. Tian T, Qiu Z, Jiang Y, Zhu D, Zhou X. Exploiting the orthogonal CRISPR-Cas12a/Cas13a trans-cleavage for dual-gene virus detection using a handheld device. Biosens Bioelectron. 2022. Vol. 196:113701

136. Wang P, Ma C, Zhang X, Chen L, Yi L, Liu X, et al.. A Ligation/recombinase polymerase amplification assay for rapid detection of SARS-CoV-2. Front Cell Infect Microbiol. 2021. Vol. 11:680728

137. Choi MH, Lee J, Seo YJ. Combined recombinase polymerase amplification/rkDNA-graphene oxide probing system for detection of SARS-CoV-2. Anal Chim Acta. 2021. Vol. 1158:338390