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Chapter 4.1. Cholera diagnosis: current and future technologies for point-of-care diagnosis in less developed countries.

Authors: 

Engku Nur Syafirah Engku Abd Rahman, Nurul Najian Aminuddin Baki and Chan Yean Yean

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Carsten Stormer
After ceasefire, many barriers to access remain,
 including security threats for health care workers 
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When you're working on development issues, optimism
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Jim Yong Kim

Introduction

Despite progress in medicine and biomedical science technologies, cholera remains an epidemic and endemic disease in many regions worldwide, especially in developing areas that currently lack effective sanitation. A cholera epidemic started in the Republic of Haiti in October 2010. By October 17, 2013, the Ministère de la Santé Publique et de la Population reported 684,085 occurrences with 8,361 deaths since the start of the outbreak, and 380,846 (55.4%) patients were admitted to hospital [1].

Vibrio cholerae is mainly transmitted by consuming contaminated food or water, and/or via the fecal–oral route. The characteristic watery diarrhea is caused by a cholera toxin produced by toxigenic Vibrio strains [2, 3], and symptoms such as severe dehydration and abdominal cramps are caused by rapid and extreme fluid electrolyte imbalances [4]. Toxigenic V. cholerae are currently known to belong to two existing serogroups: O1 and O139 Bengal, based on serological classification of the heat-stable surface somatic O antigen [2, 5]. Toxigenic serogroup O1 Vibrio has been responsible for a series of catastrophic cholera pandemics. Massive Asiatic cholera pandemics were stimulated by the Classical and El Tor biotypes from V. cholerae serogroup O1 strains, and extensive epidemics in Bangladesh and India were caused by O139 Bengal strains. The current (at the time of writing) seventh cholera pandemic is associated with V. cholerae serogroup O1 El Tor [6], and the rapid spread of the O139 serogroup in Southeast Asia from 1992 has aroused fears that it might lead to a future pandemic [7]. In contrast to the toxigenic serogroups O1 and O139, non-O1/non-O139 are considered to be non-toxigenic because of the lack of agglutination with O antiserum in biochemical tests [6].

The main pathogenic features of cholera are well established. When a person ingests contaminated water or food containing toxigenic V. cholerae, the bacteria colonize the small intestine using toxin-co-regulated pili and react with the natural receptor, monosialoganglioside GM1, on the intestinal epithelium. Once attached, the bacteria release cholera enterotoxin into the extracellular environment, thereby altering the regulation of cyclic adenosine monophosphate (cAMP) synthesis and subsequently disrupting ion transport in the intestinal epithelial cells [8]. This results in the release of electrolytes and water into the intestine, together with the bacteria. Cholera enterotoxin, or choleragen, is expressed by ctxA and ctxB genes, both of which are significant causative factors in cholera pathogenesis.

Although cholera is curable with oral rehydration therapies involving fluid and electrolyte replacement, in addition to antibiotics, it remains an extremely debilitating and potentially fatal disease in developing countries with inadequate sewage systems and clean water supplies. Cholera investigations in the clinical laboratory have focused mainly on conventional diagnostic methods; the gold standard method is culture and identification of the etiologic agent by biochemical examination. However, these tests are time-consuming in the case of bacterial culture. Moreover, they are tedious and costly, and require skilled personnel. Furthermore, poorly equipped small research laboratories in undeveloped and developing countries may lack the facilities to isolate and detect this organism.

Recent studies have therefore attempted to develop a specific diagnostic assay to allow the rapid detection of V. cholerae, thereby enabling timely clinical intervention. Such techniques might be further developed into point-of-care (POC) diagnostic systems, thereby raising the performance level to meet the ASSURED criteria of an ideal diagnosis technique, i.e., Affordable, Sensitive, Specific, User-friendly, Rapid, Equipment-free, and Delivered [9]. In this paper, we review the current diagnostic techniques (Figure 1) based on the detection of the cholera bacterium or cholera toxin, and consider the anticipated technologies for POC diagnostics in the developing world.

Figure 1. Summary of conventional methods to the latest technology for diagnosis of V. cholerae

Cholera diagnosis: from the conventional method to the latest technology

Isolation and identification of V. cholerae bacterium

Stool samples are collected from suspected cholera patients prior to antibiotic therapy. The samples are collected in suitable transport media such as Cary Blair to ensure good recovery of the bacteria. Alkaline peptone water (APW) is used as an enriched medium, and thiosulfate-citrate-bile salts-sucrose agar (TCBS) is commonly chosen as a commercially available selection medium [10]. These media are utilized whenever there is suspected asymptomatic contagion, in convalescent patients, and whenever there are likely to be increased numbers of competing organisms present. However, during outbreaks, collected liquid feces samples contain high concentrations of V. cholerae (107–108/mL), and an enrichment medium is not needed. In formed stool samples, however, APW medium (pH 8.4–9.2) is commonly used to recover the bacteria after a 6–8-hour incubation period [4]. Other less commonly used enrichment media include APW with tellurite broth, sodium-gelatin phosphate broth [11], and Monsur’s tellurite-taurocholate broth [12].

V. cholerae belongs to the family Vibrionaceae. It is an anaerobic, non-spore-forming, gram-negative rod, with respiratory and fermentative metabolism capabilities. It produces a positive result in the oxidase biochemical test, is motile via a single, sheathed flagellum, and is able to reduce nitrate [2, 4]. Freshly-cultured Vibrio bacteria in APW are inoculated on TCBS agar and incubated overnight or for 24 hours at 35–37°C. V. cholerae develop as large, yellow, shiny, smooth colonies 2–4 mm in diameter on agar plates. The yellow color is caused by the fermentation of sucrose; in contrast, sucrose-non-fermenting organisms like Vibrio parahemolyticus produce green or blue-green colonies on TCBS. V. cholerae O1 serogroup growth can be inhibited by O/129 compound (2,4-diamino-6,7-diisopropylpteridine), but resistance to this chemical is currently increasing in connection with trimethoprim-sulfamethoxazole resistance [2, 13, 14]. However, V. cholerae O139 isolates are resistant to O/129 compound [15].

Biochemical tests

Biochemical tests prior to testing with O1 and O139 antisera are generally considered unnecessary, though some screening tests may be beneficial in the event of limited serum supplies. The oxidase test is crucial for distinguishing between V. cholerae, which is oxidase-positive, and Enterobacteriaceae, which are all oxidase-negative. Additional biochemical screening tests, such as the String test [16] triple sugar iron agar [17], the Gram stain, and wet mount using dark-field and phase-contrast microscopy to check bacterial motility can be used before testing with antisera [10].

Serological tests

Agglutination tests using polyvalent antisera raised against the O1 and O139 antigens are used to confirm the identities of V. cholerae O1 and O139 serogroups, respectively. Freshly cultured V. cholerae on non-selective agar is used to avoid false-negative results, which can be produced by freshly cultured bacteria on TCBS agar [18]. Agglutination with polyvalent antisera against the O1 antigen confirms the O1 strain of V. cholerae, whereas agglutination with polyvalent antisera against O139 antiserum confirms the O139 strain. However, in the event of agglutination with O1 and O139 antisera, likely V. cholerae O1 serogroup samples should be screened with Inaba and Ogawa sera, because a positive reaction with either of these is confirmation of the V. cholerae O1 serogroup. The V. cholerae O1 serogroup can be separated into three serotypes: Ogawa, Inaba, and Hikojima, which can be identified on the basis of agglutinating activity in type-specific O antigen monovalent sera [5]. The O antigen in the O1 serogroup exhibits different subtypes indicated by the factors A, B, and C. The A factor is believed to be a D-perosamine homopolymer, whereas factors B and C are still unknown, and the differences between these three subtypes are largely quantitative. For example, the Ogawa strains possess A and B factors and a minor portion of C factor, whereas the Inaba subtype only possesses the A and C factors. The Hikojima strain raises all three antigens and thus reacts with both Inaba and Ogawa sera; however, this strain is rare and unstable [6].

Determination of V. cholerae O1 biotypes may be unnecessary for patient intervention, but the collected data are significant for identifying the infection source in epidemiological- and public health-related terms, especially when initial suspected cholera samples are isolated and collected in certain geographic areas. Historically, V. cholerae O1 serogroups consist of two biotypes, namely the Classical and El Tor biotypes [19, 20]. Isolates from the sixth pandemic were mainly of the Classical biotype, but this was replaced by the El Tor biotype associated with sporadic diarrhea in the seventh pandemic, when it was first isolated at a quarantine site in Sinai (1905) [21]. However, isolation of the El Tor biotype has increased, and it is now the predominant biotype worldwide. The Voges–Proskauer test is normally applied to differentiate between these two biotypes [19]. El Tor isolates generally produce positive results, whereas Classical isolates generally give negative results because of loss of function and lack of hemolysin associated with an 11-base deletion in the hemolysin structural gene [8, 22]. Regarding identification of the V. cholerae O139 serogroup, if bacterial identification using the specific antiserum fails, checking for the presence of choleragen and confirmation of the O139 antigen can be performed. No biotypes or serotypes are identified in the O139 serogroup. V. cholerae O139 strains are genetically similar to V. cholerae O1 strains from the seventh cholera pandemic [23-25], with the 22-kb O1 O antigen gene cluster replaced by the 35-kb O139 gene cluster via recombination [26, 27]. The antisera mentioned above are commercially available and are applied in slide agglutination trials.

Toxin assays

The identification of V. cholerae involves the crucial distinction between strains that can and cannot produce cholera toxin. Toxin assays are not routinely performed in the diagnostic laboratory, but should be carried out during the early stages of a cholera outbreak or at a low incidence of cholera cases to confirm the pathogenicity of suspected bacteria. Cholera toxin can be detected in three ways: bioassays, immunoassays, and/or DNA-based assays [10]. Bioassays commonly involve animal studies, e.g., rabbit ileal loops are injected with cholera toxin to ascertain the volume of fluid accumulation induced by enterotoxin, and tissue culture techniques, e.g., using Y1 mouse adrenal (Y1) [28] and Chinese hamster ovary (CHO) [29] cells. The presence of cholera toxin causes morphological changes in the cells (CHO cells stretch, whereas Y1 cells become rounded), as seen under a microscope. Immunoassays such as enzyme-linked immunosorbent assays (ELISAs) and the latex agglutination test (VET-RPLA; Denka Seiken, Tokyo, Japan) are usually carried out in the laboratory. The discovery of the natural receptor of cholera toxin, GM1 ganglioside, in the human intestine has contributed to the establishment of a ganglioside-capture ELISA (GM1 ELISA) [30]. The latex agglutination technique requires high-quality cholera toxin antiserum, which binds to latex particles. Almeida et al. (1990) reported that latex agglutination provided a rapid and simple test for recognizing cholera toxin with good sensitivity and specificity, compared with ELISA (0.97 and 1.00, respectively) [31]. However, compared with DNA-based assays such as the polymerase chain reaction (PCR), these methods still have relatively low sensitivity; (VET-RPLA) and the GM1 ELISA method require about 2–3 days to produce results [32, 33].

DNA probes and PCR

Molecular-based tests are not routinely employed for V. cholerae detection and identification in the clinical laboratory, because of the ease of standard techniques. However, DNA probes provide a specific and sensitive method of detecting cholera toxin-producing bacteria, and for distinguishing between strains without cholera toxin-producing genes. Cloned DNA, restriction enzyme fragments from copied DNA, and PCR amplicons are generally used as probes. Labeling with enzymes, radioisotopes, or ligands is followed by hybridization with V. cholerae complementary DNA (cDNA) as probes; non-hybridized probes are washed out, whereas the hybridized probe can be detected using assays. However, oligonucleotide probes may be too specific when used under high-stringency conditions, which may result in the hybridization of unrelated sequences and the omission of variable sequences in the target genes. Yoh et al. (1993) reported the detection of cholera toxin using an enzyme-labeled oligonucleotide probe, but although this assay is suitable for use in clinical laboratories that use radioisotope-labeled DNA fragment probes, its low sensitivity makes it impractical for use in the field. It is therefore preferable to target relatively conserved sequences to overcome this problem, and similar arguments can be applied to PCR. This is particularly relevant in the case of assaying environmental samples of V. cholerae [34], or when working in areas of low cholera endemicity, given that the majority of environmental Vibrio O1 strains outside the indigenous areas do not carry toxigenic genes [35].

PCR is frequently used to detect cholera toxin-encoding ctx genes, which are highly conserved among V. cholerae strains. DNA amplification of ctx sequences is carried out using Taq DNA polymerase and specific primers, and the amplification products are observed by gel electrophoresis followed by staining with ethidium bromide or DNA probes. PCR is preferable to the use of DNA probes because it can rapidly amplify target sequences to a detectable level, and it only requires a small sample. Apart from conventional PCR, more sophisticated approaches exist for V. cholerae detection, such as multiplex PCR to detect O1 and O139 Vibrio strains [36], and real-time PCR assays [37], which provide more rapid V. cholerae detection than conventional PCR. However, these high-technology DNA-based tests require trained and skilled personnel to perform the task, an expensive thermocycler to run the PCR, and a fluorescence detector in the case of real-time PCR. Moreover, the facility to perform gel electrophoresis to detect the amplified products is restricted in many laboratories.

Lateral-flow dipstick (LFD) assay

Immunochromatographic assays, also called strip tests or dipstick assays, have been available for several years. The one-step test technology is a continuation of latex agglutinating activity tests, and was developed from the method described by Plotz and Singer (1956) [38]. Immunochromatographic assays are user-friendly, rapid, inexpensive in terms of production and development, and show long-term consistency in broad climatic areas. Therefore, they are potentially ideal for rapid POC testing, home-based testing, and on-site testing of environmental, clinical, food, and agricultural samples [39-43]. The LFD assay provides a reliable testing method that is probably not available in undeveloped and developing countries. Pregnancy tests, strep throat, human immunodeficiency virus, and chlamydia detection tests are examples of other common LFD tests available on the market [43].

LFD assays are based on the immunochromatography principle, and consist of four components: a specimen-loading pad, a conjugate pad comprising conjugated colloidal gold, a detection membrane consisting of a test line and control line, and an absorbent pad [43, 44]. In practice, when the sample pad of the dipstick is dipped into the sample, the aqueous sample starts to migrate towards the absorbent pad by passing through the conjugate pad and the detection membrane by capillary action. When the sample reaches the conjugate pad, the analytes within the sample bind to the colloidal gold conjugate to form an analyte–antibody binding complex. The analyte–antibody complexed with colloidal gold then moves towards the capture target, which is immobilized on the membrane, to produce a distinct red line on the detection membrane in the case of a positive reaction, or no red line in the case of a negative result. A control line, produced by trapping excess gold colloidal conjugate, indicates that the test is complete and, importantly, that the results are valid [43, 45]. The red signal from the colloidal gold can be read qualitatively within 10–15 min, and the intensity of the line represents the amount of analyte present in the sample. The sandwich method and competitive method are the two main methods used in the LFD assay [46], depending on the purpose of the assay. The sandwich method is commonly used to detect larger analytes with multiple binding sites, such as human chorionic gonadotropin in pregnancy tests, whereas the competitive method is often applied to smaller analytes with a single antigenic determinant [43].

Some studies have reported the detection of V. cholerae using LFD assays, either by immunochromatographic [42, 47] or nucleic acid-based techniques [39, 40, 48]. A single-step LFD has been developed for the simultaneous detection of V. cholerae O1 and O139 serogroups using specific monoclonal antibodies against lipopolysaccharides conjugated with colloidal gold nanoparticles [42]. This assay has the ability to detect as few as 108 and 107 colony-forming units (CFU)/mL V. cholerae O1 and O139, respectively, with overall sensitivity for O1 of 93–100% and specificity for O1 of 100%, compared with the O1 dipstick test, which showed a sensitivity of 94.2% and a specificity of 98% [47]. The sensitivity and specificity for O139 were 100% and 98–100%, respectively, compared with the 100% sensitivity and 92.5% specificity, respectively, demonstrated by Nato et al. (2003). DNA-based LFD assays for detection of the ctxA gene PCR amplicons [39] provide better specificity by using target sequence-specific complementary sequences, with the potential for rapid and specific recognition of V. cholerae during outbreaks; however, the need for a PCR thermocycler probably restricts the use of this technique in resource-limited settings. The nucleic acid LFD biosensor test has a limit of detection of 0.3 ng, which is less than the 5 ng reported by Chua et al. [40], and the 8 ng reported by BlaĆŸková et al. [48], and was also able to detect stool-spiked clinical samples with 100% sensitivity and specificity. Moreover, the dry reagent lateral flow produced by the cold-chain-free feature during transfer and storage can accelerate diagnosis during outbreaks in limited-resource areas [39, 42].

Loop-mediated isothermal amplification (LAMP)

LAMP is a widely recognized and effective emerging DNA amplification technique that is gaining popularity among researchers for the early detection of microbial infections because it is rapid, simple, accurate, and cost-effective [49-55]. As originally reported by Notomi et al. (2000), LAMP is a fast and unsophisticated one-step process that amplifies specific DNA target sequences under isothermal conditions in under 1 hour. It requires a Bst DNA polymerase with strand displacement activity and three sets of specifically designed primers, and all the reagents are incubated together in one tube. LAMP amplicons can be visualized with the naked eye, either by prior amplification with calcein [55], hydroxynaphthol blue [56-58], or SYBR Green I [59], by prior detection by visualizing turbidity [60], and/or by including SYBR Green I or ethidium bromide [61] after amplification. Agarose gel electrophoresis [55, 61, 62] and real-time monitoring of gene amplification using a turbidimeter [63] can also be employed. LAMP uses a heating block or a water bath, rather than the thermal cycler required for PCR.

The application of LAMP for detecting various pathogens has been widely studied, and several commercial LAMP diagnostic kits exist for detecting Salmonella, Escherichia coli, Campylobacter, and Legionella [64]. Early studies have detected V. cholerae using the LAMP method [63], and have demonstrated that LAMP provides a fast and sensitive means of detecting toxigenic Vibrio strains compared with standard biochemical methods and PCR tests. LAMP accurately recognized 34 toxigenic Vibrio strains, but did not recognize 13 non-cholera toxin-producing V. cholerae strains or 53 non-Vibrio bacterial strains. The LAMP test was 10-fold more sensitive than conventional PCR, with 7.8 x 102 CFU/g (1.4 CFU per reaction) when tested on cholera toxin-producing Vibrio in spiked human feces, using detection by real-time turbidimetry [63]. A LAMP study was also carried out on Vibrio parahaemolyticus, a seafood-borne infectious agent, and the sensitivity of the assay on spiked shrimp specimens was 5.3 x 102 per mL/g (2.0 CFU per reaction), which was 10-fold more sensitive than standard PCR [65]. Its simplicity, rapidity, sensitivity, and cost-effectiveness suggest that the LAMP technique is suitable for the rapid diagnosis of contagious illnesses, is relevant for on-site analysis, and has the potential to provide a genetic POC assay to eliminate or confirm the presence of communicable diseases in undeveloped and developing countries.

Biosensors

A biosensor is defined as an analytical instrument that includes a biologically related constituent and a physical electronic transducer that detects, transmits, and records information on physiological and biochemical changes to produce a measurable electrical response that is proportional to the analyte concentration [66-68]. Generally, the biological identification component of the biosensor reacts to the target, and the transducer then transforms the biological reactions to a noticeable signal, which is evaluated as an electrochemical, optical, acoustic, electronic, mechanical, or calorimetric signal that is correlated with the concentration of the analyte [66, 67]. Biosensors have been comprehensively studied and intensively employed in numerous applications since they were first developed for glucose detection by Clark and Lyon in 1962. They have been used in applications ranging from environmental monitoring and public health to food safety, pathogenic bacteria detection, analytical chemistry, agricultural industries, and homeland security [69-71]. Numerous biological elements have been used in the fabrication of biosensors, including enzymes, tissues, antibodies, microorganisms, cofactors, and even the cells of higher organisms. Among these, enzymes are extensively used as biological elements because of the high sensitivity and specificity of their activities [66]; however, enzyme purification is time-consuming and costly, and alternatives are therefore needed. Microorganisms such as bacteria, yeasts, and algae can offer an alternative, and have the advantages of mass production, easy genetic manipulation via mutation [66], and better viability and stability in vitro [72], which enhance biosensor performance [67].

The development of microbial biosensors involves the use of suitable transducers according to the purpose of the biosensor, where the measured signal detected by the transducer can be correlated with the analyte concentration. The establishment of cost-effective and disposable biosensors for on-site clinical and environmental diagnoses, especially electrochemical immunosensors based on amperometric and voltammetric methods, has recently attracted the attention of researchers [73, 74]. Several platforms have been reported in the case of V. cholerae detection, including electrochemical [73-75], surface plasmon resonance (SPR) [76], quartz crystal microbalance [77, 78], and microcantilever [79] methods, the cost-effectiveness and simplicity of operation of which offer the potential for POC diagnostics. Electrochemical immunosensors are based on the high specificity of conventional immunochemical methods together with the low detection limit of advanced electrochemical systems [73]. Electrodes for electrochemical immunosensors can be constructed specifically for field applications. Numerous types of electrodes can be used, including carbon paste [80, 81], gold [78, 82], platinum [83], glassy carbon [70], and the increasingly popular screen-printed electrodes (SPEs) [84, 85]. SPEs are used extensively because they are inexpensive, can be mass-produced [86], reproduced [86, 87], and used singly [88].

Compared with immunological-based detection methods, it has been claimed that DNA-based detection is more sensitive and specific for the diagnosis of particular pathogenic bacteria. Furthermore, DNA probe-based electrochemical biosensors have also been reported for the detection of V. cholerae using carbon SPEs [89]. Probe-based electrochemical biosensors produced for chronoamperometric detection show up to 100% specificity for the single-stranded asymmetric lolB gene PCR product, with a sensitivity limit of detection as low as 0.85 ng/µL O139 strain of genomic V. cholerae DNA [75]. Implementation of the sandwich-type double hybridization technique used in this study avoided the non-specific amplifications that can be produced by electrophoretic analysis (caused by primers mispriming target DNA sequences) [90], as a result of the selective utilization of both capture and detection probes selectively attached to the single-stranded DNA of the PCR amplicon [75].

Immunosensors based on SPR quantify antigens using antibody particles immobilized on the sensor surface, which are able to detect target analytes in composite biological media with high sensitivity and specificity [91]. A surface plasmon is a quantum of plasma oscillation that spreads at the metal–dielectric surface. The loose electron gas from the metal surface is driven in a discrete mode by the external laser field, which subsequently causes the spatial distribution of charges and produces an electrical field at the metal–dielectric interface. SPR is thus exceedingly sensitive to the interfacial architecture. Shifts in plasmon resonance occur as a result of adsorption, and can thus be used to track surface changes with high accuracy [92]. Jyoung et al. (2006) developed an SPR immunosensor for V. cholerae O1 strain detection involving construction of a layer of protein G on a mixed self-assembled monolayer consisting of 11-mercaptoundecanoic acid and hexanethiol for immobilization of the antibody. Examination of the constructed layer by SPR spectroscopy permits the observation of plasmon resonance angle shifts. The immunosensor showed specific binding capacity and a detection range of 105–109 cells/mL [76]. Given that the concentration of V. cholerae in feces during outbreaks is usually around 106 CFU/mL [93], the SPR immunosensor is appropriate for the identification of V. cholerae in fecal specimens.

Alfonta et al. (2001) developed an electrochemical choleragen-detection method based on the transduction of a microgravimetric quartz crystal microbalance. They used horseradish peroxidase (HRP) and GM1-functionalized liposomes as the detection elements for amplified choleragen detection based on the highly specific detection of cholera toxin by the ganglioside GM1. A monoclonal antibody against the B subunit of cholera toxin associated with protein G is set up as a single layer on a gold electrode or a gold/quartz crystal for the sensing interface. Electronic transducers work by the “sandwich-type” concept, which detects cholera toxin attached to the anti-cholera toxin antibody, and the HRP-GM1-ganglioside-functionalized-liposome precipitates the insoluble product 3 (produced by oxidation of 4-chloronaphthol (2) in the presence of H2O2) on the gold/quartz crystal transducers to amplify the sensing result. The technique has been used to detect cholera toxin with a sensitivity of 1.0 × 10-13 M [77].

Biosensor technology integrated with a microcantilever platform can produce a powerful biosensing design with excellent sensitivity and a low detection limit [94, 95]. Combined with dynamic force microscopy (DFM) and a microcantilever in an atomic force microscope (AFM), the concentration of V. cholerae O1 strain monoclonal antibody adsorbed on the gold-coated microcantilever surface by the self-assembly monolayer technique can be detected and measured based on the resonance frequency shift data. In DFM, a piezoelectric actuator drives the microcantilever at its resonance frequency, and whenever the target particles (V. cholerae O1 strain antibody) are absorbed onto the microcantilever, its resonance frequency is reduced because of the increased mass [79, 96]. Changes in the mass of the microcantilever thus create a resonance frequency shift relative to the quantity of adsorbed particles [96]. Sungkanak et al. (2010) reported a high sensitivity of about 146.5 pg/Hz using this technology, with a limit of detection as low as approximately 1 × 103 CFU/mL, which is lower than the detection limits for ELISA and amperometric immunosensors [73]. The technique is therefore suitable for use in general microbiology laboratories for the detection of V. cholerae in food production, without the need for pre-enrichment procedures [79].

Future prospects for POC diagnosis

The discovery and establishment of nucleic acid technology systems such as PCR and LAMP [49, 53], and biosensors [97, 98] re useful for the development of POC diagnostic devices or bedside testing instruments that could eliminate and/or confirm the presence of communicable contagions, especially cholera, in poor, undeveloped, and developing countries. The worldwide chaos caused by the cholera pandemics in the 1990s, together with the unexpected reappearance of epidemic cholera from the O1 El Tor strain of V. cholerae in Latin America after a century of absence in the [99] and the appearance of a new V. cholerae O139 strain in late 1992 in Southern India [100], have highlighted the problems of this transmissible infectious disease in these poor regions. To prevent such epidemics spreading to non-endemic areas, it is essential to develop a diagnostic device that is widely applicable, especially in settings with limited resources, and which can be used at any time and by anyone, including nurses and non-skilled personnel, to screen infected and suspected cholera patients.

New technologies have been developed and studied, some of which have been applied to various fields of analytical science. Micro-total analysis systems (µTAS), popularly known as lab-on-a-chip (LOC), represent a new technology concept developed in 1979 by Standford University researchers based on gas chromatography [101]. LOC technology resembles a miniaturized analytical laboratory that allows all the conventional analytical steps practiced in the laboratory, including pre-treatment of the sample, sample separation, signal amplification, and signal recognition, to be carried out on a small chip device. This analysis is fast, efficacious, and automatic [60, 102]. Its small size, low sample volume, and rapidity make it one of the most powerful applications for use in remote settings as a potential POC device [103]. Nucleic acid technologies generally involve labor-intensive procedures, extraction of samples to obtain pure genomic DNA, amplification of the specific DNA target, and recognition of the desired products, and therefore present a promising application for LOC technology.

Whereas the LOC concept has been applied to PCR [104], methods such as LAMP have also been integrated into eight-channel microfluidic chips, the results of which can be read either by the naked eye as a result of the formation of an insoluble byproduct, or via fiber optic sensors that measure turbidity absorbance. Known as micro-LAMP (µLAMP), this technology was used to detect pseudorabies virus (PRV) by Fang et al. (2010), who claimed that µLAMP was straightforward in terms of manipulation and was able to quantitatively analyze DNA samples with high sensitivity and specificity using a sample volume of only 0.4 µL, with completion of the reaction within 1 hour at 63°C. The assay’s limit of detection is 10 fg of target sequence, which is 100–1000-fold more sensitive than conventional PCR for the diagnosis of PRV, with 100% specificity [105]. In 2012, the same researchers developed a portable integrated LAMP microchip (iµLAMP) that provides rapid DNA release, exponential signal amplification, and a readout that is detectable by the naked eye, either in single or multiplex format, for the identification of bacteria [106]. Apart from the LAMP method, several isothermal techniques have also been established as miniaturized systems, such as helicase-dependent amplification, nucleic acid sequence-based amplification, strand displacement amplification, and rolling circle amplification [102], which are suitable for the detection of various pathogens, especially V. cholerae. Moreover, several novel isothermal techniques such as exponential amplification reaction, signal-mediated amplification of RNA technology, and isothermal and chimeric primer-initiated amplification of nucleic acids have also been described by Asiello and Baeumner (2011).

The combination of LAMP and microfluidics shows great promise in terms of the development of easy-to-use, portable, and disposable LOC devices for the detection of V. cholerae on-site, especially during massive outbreaks. Such devices have the potential to produce multiple, high-throughput results and have several advantages over PCR: they do not require thermal cycling, their energy consumption is low, and they are portable. The need for only a small sample volume makes the methods cost-effective, which must be considered when proposing a microsystem for use in limited-resource settings. A biosensor and microfluidics combination has also been developed that only requires a short processing time, including sample incubation [98], and a small sample volume, and which eliminates extensive cell culture processes [97]. These anticipated technologies have the potential to provide almost perfect LOC-based systems using nucleic acid technology, which would facilitate the detection of amplicons via their simplicity, effectiveness, rapidity, and cost-effectiveness in the near future.

Conclusions

In this review, we discussed the current and anticipated technologies for cholera diagnosis, with the aim of developing POC diagnosis devices for use in undeveloped and developing countries. New technologies for cholera diagnosis need to demonstrate fast, cost-efficient, and accurate diagnoses based on sensitivity and specificity criteria for cholera patients, to provide POC diagnostic tools in the future. These technologies will improve the quality of life of patients in poor and limited-resource areas where diseases are endemic, thereby highlighting the need to perfect diagnosis systems to prevent the spread of such illnesses.

References

  1. CDC, Cholera in Haiti [Online]. Available at: http://wwwnc.cdc.gov/travel/notices/watch/haiti-cholera. Centers for Disease Control and Prevention 2013.
  2. Faruque, S.M., M.J. Albert, and J.J. Mekalanos, Epidemiology, genetics, and ecology of toxigenic Vibrio cholerae. Microbiology and Molecular Biology Reviews, 1998. 62(4): p. 1301-+.
  3. Singh, D.V., et al., Molecular analysis of Vibrio cholerae O1, O139, non-O1, and non-O139 strains: clonal relationships between clinical and environmental isolates. Appl Environ Microbiol, 2001. 67(2): p. 910-21.
  4. Kaper, J.B., J.G. Morris, Jr., and M.M. Levine, Cholera. Clin Microbiol Rev, 1995. 8(1): p. 48-86.
  5. Gardner, A.D. and K.V. Venkatraman, The Antigens of the Cholera Group of Vibrios. J Hyg (Lond), 1935. 35(2): p. 262-82.
  6. Nair, G., Vibrio cholerae. In: Guidelines for drinking-water quality. World Health Organization, Geneva, 2002. 2nd Ed.
  7. Bhattacharya, S.K., et al., Clinical profile of acute diarrhoea cases infected with the new epidemic strain of Vibrio cholerae O139: designation of the disease as cholera. J Infect, 1993. 27(1): p. 11-5.
  8. Reeves, P.R. and R.T. Lan, Cholera in the 1990s. British Medical Bulletin, 1998. 54(3): p. 611-623.
  9.  Urdea, M., et al., Requirements for high impact diagnostics in the developing world. Nature, 2006. 444 Suppl 1: p. 73-9.
  10. CDC, Laboratory methods for the diagnosis of Vibrio cholerae. Centers for Disease Control and Prevention. Atlanta, Georgia, USA. , 1994.
  11. Rennels, M.B., et al., Selective Vs Nonselective Media and Direct Plating Vs Enrichment Technique in Isolation of Vibrio-Cholerae - Recommendations for Clinical Laboratories. Journal of Infectious Diseases, 1980. 142(3): p. 328-331.
  12. Monsur, K.A., Bacteriological diagnosis of cholera under field conditions. Bull World Health Organ, 1963. 28(3): p. 387-9.
  13. Nath, G. and S.C. Sanyal, Emergence of Vibrio cholerae O1 resistant to vibriostatic agent 0/129. Lancet, 1992. 340(8815): p. 366-7.
  14. Ramamurthy, T., et al., Taxonomical implications of the emergence of high frequency of occurrence of 2,4-diamino-6,7-diisopropylpteridine-resistant strains of Vibrio cholerae from clinical cases of cholera in Calcutta, India. J Clin Microbiol, 1992. 30(3): p. 742-3.
  15. Albert, M.J., et al., Large outbreak of clinical cholera due to Vibrio cholerae non-O1 in Bangladesh. Lancet, 1993. 341(8846): p. 704.
  16. Neogy, K.N. and A.C. Mukherji, A study of the "string" test in vibrio identification. Bull World Health Organ, 1970. 42(4): p. 638-40.
  17. Skillern, J.K. and T.L. Overman, Oxidase Testing from Kliglers Iron Agar and Triple Sugar Iron Agar Slants. Current Microbiology, 1983. 8(5): p. 269-271.
  18. Gangarosa, E.J., et al., Laboratory methods in cholera: isolation of Vibrio cholerae (el tor and classical) on TCBS medium in minimally equipped laboratories. Trans R Soc Trop Med Hyg, 1968. 62(5): p. 693-9.
  19. Dziejman, M., et al., Genomic characterization of non-O1, non-O139 Vibrio cholerae reveals genes for a type III secretion system. Proceedings of the National Academy of Sciences of the United States of America, 2005. 102(9): p. 3465-3470.
  20. Karaolis, D.K., R. Lan, and P.R. Reeves, The sixth and seventh cholera pandemics are due to independent clones separately derived from environmental, nontoxigenic, non-O1 Vibrio cholerae. J Bacteriol, 1995. 177(11): p. 3191-8.
  21. Osin, A.V., et al., [Comparative genomic analysis of vibrio cholerae El Tor preseventh and seventh pandemic strains isolated in various periods]. Genetika, 2005. 41(1): p. 53-62.
  22. Alm, R.A., U.H. Stroeher, and P.A. Manning, Extracellular proteins of Vibrio cholerae: nucleotide sequence of the structural gene (hlyA) for the haemolysin of the haemolytic El Tor strain 017 and characterization of the hlyA mutation in the non-haemolytic classical strain 569B. Mol Microbiol, 1988. 2(4): p. 481-8.
  23. Faruque, S.M., et al., Molecular analysis of rRNA and cholera toxin genes carried by the new epidemic strain of toxigenic Vibrio cholerae O139 synonym Bengal. J Clin Microbiol, 1994. 32(4): p. 1050-3.
  24. Karaolis, D.K., R. Lan, and P.R. Reeves, Molecular evolution of the seventh-pandemic clone of Vibrio cholerae and its relationship to other pandemic and epidemic V. cholerae isolates. J Bacteriol, 1994. 176(20): p. 6199-206.
  25. Popovic, T., et al., Molecular subtyping of toxigenic Vibrio cholerae O139 causing epidemic cholera in India and Bangladesh, 1992-1993. J Infect Dis, 1995. 171(1): p. 122-7.
  26. Comstock, L.E., et al., Cloning and sequence of a region encoding a surface polysaccharide of Vibrio cholerae O139 and characterization of the insertion site in the chromosome of Vibrio cholerae O1. Mol Microbiol, 1996. 19(4): p. 815-26.
  27. Mooi, F.R. and E.M. Bik, The evolution of epidemic Vibrio cholerae strains. Trends Microbiol, 1997. 5(4): p. 161-5.
  28. Donta, S., Differentiation between the steroidogenic effects of cholera enterotoxin and adrenocorticotropin through use of a mutant adrenal cell line. J Infect Dis, 1974. 129: p. 728-731.
  29. Guerrant, R.L., et al., Cyclic adenosine monophosphate and alteration of Chinese hamster ovary cell morphology: a rapid, sensitive in vitro assay for the enterotoxins of Vibrio cholerae and Escherichia coli. Infect Immun, 1974. 10(2): p. 320-7.
  30. Svennerholm, A.M. and J. Holmgren, Identification of Escherichia-Coli Heat-Labile Enterotoxin by Means of a Ganglioside Immunosorbent Assay (Gm1-Elisa) Procedure. Current Microbiology, 1978. 1(1): p. 19-23.
  31. Almeida, R.J., et al., Comparison of a latex agglutination assay and an enzyme-linked immunosorbent assay for detecting cholera toxin. J Clin Microbiol, 1990. 28(1): p. 128-30.
  32. Varela, P., et al., Direct detection of Vibrio cholerae in stool samples. J Clin Microbiol, 1994. 32(5): p. 1246-8.
  33. Yoh, M., et al., Development of an Enzyme-Labeled Oligonucleotide Probe for the Cholera-Toxin Gene. J Clin Microbiol, 1993. 31(5): p. 1312-1314.
  34. Wright, A.C., et al., Development and testing of a nonradioactive DNA oligonucleotide probe that is specific for Vibrio cholerae cholera toxin. J Clin Microbiol, 1992. 30(9): p. 2302-6.
  35. Minami, A., et al., Cholera Enterotoxin Production in Vibrio-Cholerae-O1 Strains Isolated from the Environment and from Humans in Japan. Appl Environ Microbiol, 1991. 57(8): p. 2152-2157.
  36. Hoshino, K., et al., Development and evaluation of a multiplex PCR assay for rapid detection of toxigenic Vibrio cholerae O1 and O139. Fems Immunology and Medical Microbiology, 1998. 20(3): p. 201-207.
  37. Fykse, E.M., et al., Detection of Vibrio cholerae by real-time nucleic acid sequence-based amplification. Appl Environ Microbiol, 2007. 73(5): p. 1457-66.
  38. Plotz, C.M. and J.M. Singer, The latex fixation test. I. Application to the serologic diagnosis of rheumatoid arthritis. Am J Med, 1956. 21(6): p. 888-92.
  39. Ang, G.Y., C.Y. Yu, and C.Y. Yean, Ambient temperature detection of PCR amplicons with a novel sequence-specific nucleic acid lateral flow biosensor. Biosens Bioelectron, 2012. 38(1): p. 151-6.
  40. Chua, A., et al., A rapid DNA biosensor for the molecular diagnosis of infectious disease. Biosens Bioelectron, 2011. 26(9): p. 3825-31.
  41. Kumanan, V., et al., A biosensor assay for the detection of Mycobacterium avium subsp. paratuberculosis in fecal samples. J Vet Sci, 2009. 10(1): p. 35-42.
  42. Yu, C.Y., et al., Dry-reagent gold nanoparticle-based lateral flow biosensor for the simultaneous detection of Vibrio cholerae serogroups O1 and O139. Journal of Microbiological Methods, 2011. 86(3): p. 277-282.
  43. Zhang, G., J. Guo, and X. Wang, Immunochromatographic lateral flow strip tests. Methods Mol Biol, 2009. 504: p. 169-83.
  44.  Kumar, R. and R. Sinha, Colloidal gold based dipstick strip for detection of genetically modified crops and produce. Int J Pharma Bio Sci, 2011. 2: p. 110.
  45. Chandler, J., T. Gurmin, and N. Robinson, The place of gold in rapid tests. IVD Technol, 2000. 6: p. 37-49.
  46. Weiss, A., Concurrent engineering for lateral-flow diagnostics. IVD Technol, 1999. 5: p. 48-57.
  47. Nato, F., et al., One-step immunochromatographic dipstick tests for rapid detection of Vibrio cholerae O1 and O139 in stool samples. Clin Diagn Lab Immunol, 2003. 10(3): p. 476-8.
  48. Blazkova, M., et al., Immunochromatographic strip test for detection of genus Cronobacter. Biosens Bioelectron, 2011. 26(6): p. 2828-34.
  49. Dhama, K., et al., Loop-mediated isothermal amplification of DNA (LAMP): a new diagnostic tool lights the world of diagnosis of animal and human pathogens: a review. Pak J Biol Sci, 2014. 17(2): p. 151-66.
  50. Focke, F., I. Haase, and M. Fischer, Loop-mediated isothermal amplification (LAMP): methods for plant species identification in food. J Agric Food Chem, 2013. 61(12): p. 2943-9.
  51. Notomi, T., et al., Loop-mediated isothermal amplification of DNA. Nucleic Acids Res, 2000. 28(12): p. E63.
  52. Okada, K., et al., A rapid, simple, and sensitive loop-mediated isothermal amplification method to detect toxigenic Vibrio cholerae in rectal swab samples. Diagn Microbiol Infect Dis, 2010. 66(2): p. 135-9.
  53. Parida, M., et al., Loop mediated isothermal amplification (LAMP): a new generation of innovative gene amplification technique; perspectives in clinical diagnosis of infectious diseases. Rev Med Virol, 2008. 18(6): p. 407-21.
  54. Sidoti, F., et al., Alternative Molecular Tests for Virological Diagnosis. Molecular Biotechnology, 2013. 53(3): p. 352-362.
  55. Tomita, N., et al., Loop-mediated isothermal amplification (LAMP) of gene sequences and simple visual detection of products. Nature Protocols, 2008. 3(5): p. 877-882.
  56. Dai, T.T., et al., Development of a loop-mediated isothermal amplification assay for detection of Phytophthora sojae. FEMS Microbiol Lett, 2012. 334(1): p. 27-34.
  57. Goto, M., et al., Colorimetric detection of loop-mediated isothermal amplification reaction by using hydroxy naphthol blue. Biotechniques, 2009. 46(3): p. 167-72.
  58. Zhang, Y.W., et al., Development of loop-mediated isothermal amplification method for visualization detection of the highly virulent strains of porcine reproductive and respiratory syndrome virus (PRRSV) in China. African Journal of Biotechnology, 2011. 10(61): p. 13278-13283.
  59. Parida, M., et al., Real-time reverse transcription loop-mediated isothermal amplification for rapid detection of West Nile virus. J Clin Microbiol, 2004. 42(1): p. 257-63.
  60. Mori, Y., et al., Detection of loop-mediated isothermal amplification reaction by turbidity derived from magnesium pyrophosphate formation. Biochem Biophys Res Commun, 2001. 289(1): p. 150-4.
  61. Tsai, S.M., et al., Development of a loop-mediated isothermal amplification for rapid detection of orf virus. Journal of Virological Methods, 2009. 157(2): p. 200-204.
  62. Minami, M., et al., Use of a combination of brushing technique and the loop-mediated isothermal amplification method as a novel, rapid, and safe system for detection of Helicobacter pylori. J Clin Microbiol, 2006. 44(11): p. 4032-4037.
  63. Yamazaki, W., et al., Sensitive and rapid detection of cholera toxin-producing Vibrio cholerae using a loop-mediated isothermal amplification. Bmc Microbiology, 2008. 8.
  64. Mori, Y. and T. Notomi, Loop-mediated isothermal amplification (LAMP): a rapid, accurate, and cost-effective diagnostic method for infectious diseases. J Infect Chemother, 2009. 15(2): p. 62-9.
  65. Yamazaki, W., et al., Development of a loop-mediated Isothermal amplification assay for sensitive and rapid detection of Vibrio parahaemolyticus. Bmc Microbiology, 2008. 8.
  66. D'Souza, S.F., Microbial biosensors. Biosens Bioelectron, 2001. 16(6): p. 337-53.
  67. Su, L.A., et al., Microbial biosensors: A review. Biosens Bioelectron, 2011. 26(5): p. 1788-1799.
  68. Yogeswaran, U. and S.M. Chen, A review on the electrochemical sensors and biosensors composed of nanowires as sensing material. Sensors, 2008. 8(1): p. 290-313.
  69. Amine, A., et al., Enzyme inhibition-based biosensors for food safety and environmental monitoring. Biosens Bioelectron, 2006. 21(8): p. 1405-23.
  70.  Farghali, R.A. and R.A. Ahmed, A Novel Electrochemical Sensor for Determination of Sildenafil Citrate (Viagra) in Pure Form and in Biological and Pharmaceutical Formulations. International Journal of Electrochemical Science, 2012. 7(12): p. 13008-13019.
  71. Rodriguez-Mozaz, S., M.J.L. de Alda, and D. Barcelo, Biosensors as useful tools for environmental analysis and monitoring. Analytical and Bioanalytical Chemistry, 2006. 386(4): p. 1025-1041.
  72. Byfield, M.P. and R.A. Abuknesha, Biochemical Aspects of Biosensors. Biosens Bioelectron, 1994. 9(4-5): p. 373-400.
  73. Rao, V.K., et al., Amperometric immunosensor for the detection of Vibrio cholerae O1 using disposable screen-printed electrodes. Analytical Sciences, 2006. 22(9): p. 1207-1211.
  74. Sharma, M.K., et al., Immunological biosensor for detection of Vibrio cholerae O1in environmental water samples. World Journal of Microbiology & Biotechnology, 2006. 22(11): p. 1155-1159.
  75. Low, K.F., et al., Electrochemical genosensor for specific detection of the food-borne pathogen, Vibrio cholerae. World Journal of Microbiology & Biotechnology, 2012. 28(4): p. 1699-1706.
  76. Jyoung, J.Y., et al., Immunosensor for the detection of Vibrio cholerae O1 using surface plasmon resonance. Biosens Bioelectron, 2006. 21(12): p. 2315-2319.
  77. Alfonta, L., et al., Electrochemical and quartz crystal microbalance detection of the cholera toxin employing horseradish peroxidase and GM1-functionalized liposomes. Anal Chem, 2001. 73(21): p. 5287-95.
  78. Carter, R.M., et al., Quartz crystal microbalance detection of Vibrio cholerae O139 serotype. J Immunol Methods, 1995. 187(1): p. 121-5.
  79. Sungkanak, U., et al., Ultrasensitive detection of Vibrio cholerae O1 using microcantilever-based biosensor with dynamic force microscopy. Biosens Bioelectron, 2010. 26(2): p. 784-789.
  80. FernandezSanchez, C. and A. CostaGarcia, Adsorption of immunoglobulin G on carbon paste electrodes as a basis for the development of immunoelectrochemical devices. Biosens Bioelectron, 1997. 12(5): p. 403-413.
  81. Fernandez-Sanchez, C. and A. Costa-Garcia, Inhibition of adsorbed alkaline phosphatase activity by an anti-enzyme antibody. An approach to carbon paste immunoelectrodes. Electroanalysis, 1999. 11(18): p. 1350-1354.
  82. Cheng, Q., et al., Functional lipid microstructures immobilized on a gold electrode for voltammetric biosensing of cholera toxin. Analyst, 2004. 129(4): p. 309-14.
  83. Patel, N.G., et al., Fabrication and characterization of disposable type lactate oxidase sensors for dairy products and clinical analysis. Sensors and Actuators B-Chemical, 2000. 67(1-2): p. 134-141.
  84. Schreiber, A., et al., An immunosensor based on disposable electrodes for rapid estimation of fatty acid-binding protein, an early marker of myocardial infarction. Biosens Bioelectron, 1997. 12(11): p. 1131-1137.
  85. Wang, J., P.V. Pamidi, and K.R. Rogers, Sol-gel-derived thick-film amperometric immunosensors. Anal Chem, 1998. 70(6): p. 1171-5.
  86. Alvarezicaza, M. and U. Bilitewski, Mass-Production of Biosensors. Anal Chem, 1993. 65(11): p. A525-A533.
  87. Hart, J.P. and S.A. Wring, Screen-Printed Voltammetric and Amperometric Electrochemical Sensors for Decentralized Testing. Electroanalysis, 1994. 6(8): p. 617-624.
  88. Zhang, S., G. Wright, and Y. Yang, Materials and techniques for electrochemical biosensor design and construction. Biosens Bioelectron, 2000. 15(5-6): p. 273-282.
  89. Yean, C.Y., et al., Enzyme-linked amperometric electrochemical genosensor assay for the detection of PCR amplicons on a streptavidin-treated screen-printed carbon electrode. Anal Chem, 2008. 80(8): p. 2774-2779.
  90. Dieffenbach, C.W., T.M. Lowe, and G.S. Dveksler, General concepts for PCR primer design. PCR Methods Appl, 1993. 3(3): p. S30-7.
  91. Sakai, G., et al., A surface plasmon resonance-based immunosensor for highly sensitive detection of morphine. Sensors and Actuators B-Chemical, 1998. 49(1-2): p. 5-12.
  92. Salamon, Z., H.A. Macleod, and G. Tollin, Surface plasmon resonance spectroscopy as a tool for investigating the biochemical and biophysical properties of membrane protein systems .2. Applications to biological systems. Biochimica Et Biophysica Acta-Reviews on Biomembranes, 1997. 1331(2): p. 131-152.
  93. Martinez-Govea, A., et al., Identification and strain differentiation of Vibrio cholerae by using polyclonal antibodies against outer membrane proteins. Clinical and Diagnostic Laboratory Immunology, 2001. 8(4): p. 768-771.
  94. Hansen, K.M. and T. Thundat, Microcantilever biosensors. Methods, 2005. 37(1): p. 57-64.
  95. Thundat, T., et al., Detection of Mercury-Vapor Using Resonating Microcantilevers. Applied Physics Letters, 1995. 66(13): p. 1695-1697.
  96. Lang, H.P., et al., Nanomechanics from atomic resolution to molecular recognition based on atomic force microscopy technology. Nanotechnology, 2002. 13(5): p. R29-R36.
  97. Boehm, D.A., P.A. Gottlieb, and S.Z. Hua, On-chip microfluidic biosensor for bacterial detection and identification. Sensors and Actuators B-Chemical, 2007. 126(2): p. 508-514.
  98. Zaytseva, N.V., et al., Development of a microfluidic biosensor module for pathogen detection. Lab Chip, 2005. 5(8): p. 805-811.
  99. Tauxe, R.V. and P.A. Blake, Epidemic Cholera in Latin-America. Jama-Journal of the American Medical Association, 1992. 267(10): p. 1388-1390.
  100. Ramamurthy, T., et al., Emergence of novel strain of Vibrio cholerae with epidemic potential in southern and eastern India. Lancet, 1993. 341(8846): p. 703-4.
  101. Terry, S.C., J.H. Jerman, and J.B. Angell, Gas-Chromatographic Air Analyzer Fabricated on a Silicon-Wafer. Ieee Transactions on Electron Devices, 1979. 26(12): p. 1880-1886.
  102. Asiello, P.J. and A.J. Baeumner, Miniaturized isothermal nucleic acid amplification, a review. Lab Chip, 2011. 11(8): p. 1420-30.
  103. Chin, C.D., V. Linder, and S.K. Sia, Lab-on-a-chip devices for global health: past studies and future opportunities. Lab Chip, 2007. 7(1): p. 41-57.
  104. Kopp, M.U., A.J. Mello, and A. Manz, Chemical amplification: continuous-flow PCR on a chip. Science, 1998. 280(5366): p. 1046-8.
  105. Fang, X.E., et al., Loop-Mediated Isothermal Amplification Integrated on Microfluidic Chips for Point-of-Care Quantitative Detection of Pathogens. Anal Chem, 2010. 82(7): p. 3002-3006.
  106. Fang, X.E., et al., A portable and integrated nucleic acid amplification microfluidic chip for identifying bacteria. Lab Chip, 2012. 12(8): p. 1495-1499.