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Molecular Techniques for Infectious Diseases: History
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Contributor: Akua K. Yalley ,
Selasie Ahiatrogah
, Anna A. Kafintu-Kwashie , Gloria Amegatcher ,
Diana Prah
, Akua K. Botwe , Mildred A. Adusei-Poku ,
Evangeline Obodai
, Nicholas I. Nii-Trebi

Infectious diseases significantly impact the health status of developing countries. Historically, infectious diseases of the tropics especially have received insufficient attention in worldwide public health initiatives, resulting in poor preventive and treatment options. Many molecular tests for human infections have been established since the 1980s, when polymerase chain reaction (PCR) testing was introduced. In spite of the substantial innovative advancements in PCR technology, which currently has found wide application in most viral pathogens of global concern, the development and application of molecular diagnostics, particularly in resource-limited settings, poses potential constraints.

  • molecular diagnostics
  • polymerase chain reaction
  • tropical diseases
  • infectious diseases

1. Introduction

All over the world, and especially in African countries, infectious diseases constitute a major public health challenge and thus represent one of the greatest potential barriers to achieving the third Sustainable Development Goal. This is because, collectively, they account for approximately 20% of mortality in all age groups. In the least-developed countries, they contribute to about 33% of mortality (WHO, Geneva, Switzerland, 2006).
The infectious disease burden remains alarming the world over. Approximately 15 million people die each year because of tropical infectious diseases, with most of them living in developing countries [1]. The significance of neglected tropical diseases, most of which are poverty-driven, can be underscored in this: in May 2013, the 66th World Health Assembly of the WHO adopted a resolution, WHA66.12, requiring member states to pursue and intensify measures aimed at improving the health and social well-being of affected populations [2]. Considerably, infectious diseases such as HIV/AIDS, tuberculosis, and malaria have received significant global attention, and many have already been well-documented with appreciable references [3][4][5]. However, the same cannot be said of most neglected tropical infectious diseases. It is worthy of note that tropical diseases are not limited to the tropics. Globalization and accompanying increase in international air travel for purposes including migration, tourism, and work visits to tropical regions [6] have contributed to an equally increased incidence of tropical diseases in areas such as the United States, United Kingdom, and Europe. Surveillance and measures of effective control of infectious disease pathogens therefore represent important approaches for dealing with the global spread and threat of tropical and infectious diseases [7].
Various traditional methods exist for diagnosis of most infectious disease pathogens. Table 1 presents a cross-section of these methods. However, factors that affect the concentration of pathogens in blood or blood fractions, such as latency infections, tend to render plasma concentration of pathogens such as Ebola virus, malaria parasite, human immunodeficiency virus (HIV), and tuberculosis too low to be definitively determined by methods like ELISA or blood smear. Highly sensitive techniques are therefore required that are cost effective, have fast turn-around time, and also assure reliable detection of pathogens [8]. Invariably, almost every pathogen has a nucleic acid component, which makes it possible for molecular methods to be applied for their diagnosis, monitoring, and disease study. A few examples of the traditional molecular methods include conventional PCR, real-time PCR, chromatin immunoprecipitation analysis (ChIP), nested PCR, and multiplex PCR (real-time or conventional) [9][10][11][12]. In view of their time-tested sensitivity and specificity, molecular methods offer a very reliable means of infectious disease diagnosis. The growing challenge of the tropical and infectious disease burden makes advances in molecular methods as the mainstay of infectious disease pathogen detection and control imperative.
The need for molecular diagnostics that advance clinical care and public health delivery has never been greater. Nevertheless, there are untapped opportunities that can be harnessed in shaping technologies to address current unmet needs [13]. Emerging technologies are therefore warranted that enable the detection and quantification of pathogen burden with agility, sensitivity, and simplicity. It must be acknowledged, however, that significant challenges remain with regards to the development, regulatory approval, and integration of new technologies for use in clinical diagnostics. Considerable hurdles with using some molecular methods include the fact that they are relatively expensive; require cumbersome instrumentation and reliable electricity, among others; and often require a high level of technical expertise, thus constituting a disadvantage. This underscores the need for developing point-of-care (POC) molecular diagnostic methods that may overcome some of the challenges surrounding use of traditional molecular methods.
Table 1. Traditional or non-nucleic-acid-based methods of infectious disease diagnosis.
Infectious Disease Method Description (Common Procedures) Challenges Reference
Yaws Microscopic
examination		 This method is used for the diagnosis of yaws at stage 1 and 2 using tissue samples from skin lesions. Sensitivity is low when bacterial load is low, or treponemes viability is poor. [14]
Serological
testing		 Tests include rapid plasma reagin (RPR) and Treponema pallidum particle agglutination (TPPA). Methods unable to distinguish yaws from syphilis. [14]
Buruli ulcer Microscopic
examination		 Involves direct smear or biopsy examination to detect acid-fast bacilli. Low sensitivity. [15]
Cell culture of Mycobacterium ulcerans (MU) Cell culturing to isolate viable MU for typically 9 to 16 weeks at 29–33 °C is a confirmatory test. Culturing can take months. [16]
Histopathology Analysis is done on tissue specimens in formalin stained with eosin and
hematoxylin or other stains.		 Method is expensive and does not always provide clear-cut identification. [17]
Human African trypanosomiasis Serologic
testing		 Used for screening purposes only. Reliable test available only for T.b. gambiens. [11]
Microscopic examination Used for the staging of both T.b. gambiense and T.b. rhodesiense using CSF. Very low sensitivity. [11]
Ebola Cell culture Confirms presence of Ebola virus. Visualization is done either directly by electron microscopy or indirectly by immunofluorescence microscopy. Biosafety level 4 containment is required. [18]
Antibody
detection		 Detects antibodies in serum (of some healthy individuals) usually after 3 weeks. Time taken for antibody to be detected after infection is too long. [19][20]
Onchocerciasis Microscopic
examination		 A gold standard. This is based on the detection of microfilariae in skin snips. Sensitivity of the skin snip diminishes with decreasing skin microfilaria density. [21]
Slit-lamp
examination		 Procedure involves examination of the cornea and anterior chamber of the eye. Onchocerciasis is not the only illness that may cause ocular lesions. Lesions may be seen in other infections also. [22]
Serological
testing		 The gold standard for diagnosing most common Wuchereria bancrofti cases is antigen detection. Antibody testing also exists. Has extensive antigenic cross-reactivity with other nematodes. Antibody test is unable to distinguish current from past infection. [23]
Diethylcarbamazine (DEC) Patch Test Papule formation after application of DEC to skin confirms the presence of microfilariae. Issues with sensitivity decreases after treatment with ivermectin. [24]

2. Molecular Techniques as Applied to Yaws

The bacterium Treponema pallidum subsp. pertenue (TPE) causes yaws, a severe childhood infectious disease [14]. Yaws is now known to be prevalent in 13 nations. Some of those nations include Papua New Guinea, Solomon Islands, and Ghana [25]. However, adequate reporting data are limited [26][27]. Ghana, Papua New Guinea, and the Solomon Islands in the Southwest Pacific Ocean have reported the most incidences worldwide [25]. Yaws is spread through skin-to-skin contact [28]. Spirochetal bacteria, such as the Treponema species, are responsible for a group of diseases referred to as treponematoses. These diseases and the specific causative species are: yaws, caused by Treponema pallidum pertenue; pinta, caused by Treponema carateum; bejel, caused by Treponema pallidum endemicum; and the venereal disease syphilis, caused by Treponema pallidum [29].
Conversely, with respect to diagnosing yaws, healthcare personnel in yaws-endemic nations have two fundamental problems. First, TPE, which causes yaws, shares about 99.8% of its genomic structure with T. pallidum subsp. pallidum (TPA), the causative bacterium of syphilis [30]. As a result, all of the existing serological diagnostic techniques that detect yaws also detect syphilis [31]. Over the last two decades, there has been a growing drive using PCR techniques for treponematosis investigation. Detecting treponemal DNA using PCR only needs few treponemal chromosomal copies. As an in-house test, a number of sequences have been targeted. They include tpf-1, bmp, tpp47, tmp A, and pol A, among others [32][33][34][35][36]. These tests could be used to detect treponemes from swab specimens though they are not subspecies-specific. Unlike the case in swab samples, PCR’s utility in blood samples is generally hampered by the low amount of treponemes found in blood [37][38][39].
In recent times, methods for differentiating T. pallidum subspecies have been developed. These include a real-time PCR assay, nested PCR, a combination of PCR/RFLP analysis, and sequencing analysis [40][41][42][43][44][45][46]. Since some yaws variants harbor a primer binding-site mutation, which tends to yield false-negative PCR results, molecular diagnosis in such a case should employ primers that are targeted towards highly conserved regions [43]. An emerging feature regarding proper sample handling and storage for a successful PCR reaction involves utilization of dry swabs transported at room temperature, which has been demonstrated to perform just as well in PCR as compared to swab stored in a carrier medium and subsequently transported using a cold chain [47]. This holds promise that in resource-limited settings, the use of dry swabs to cut down costs will not compromise PCR outcomes.
Despite their utility and reliability, molecular technologies such as PCR are not readily available in the field [48], in which case other techniques such as isothermal nucleic acid amplification has some relevance, as they have some advantages compared to traditional PCR. Furthermore, unlike PCR, a technique such as loop-mediated isothermal amplification (LAMP) does not necessitate thermal cycling, thereby removing the challenges associated with using a thermal cycler [49]. LAMP therefore is an ideal technique for use in developing point-of-care (POC) tests [50][51]. Indeed, a LAMP assay that can deferentially diagnose T. pallidum and H. ducreyi (targeting (pol A) gene and 16S rRNA, respectively) has recently been developed (TPHD-LAMP) with an impressive diagnostic performance of 85–92% and 85–96% sensitivity and specificity, respectively [51]. A higher sensitivity of 100% and 96% specificity of the LAMP assay (compared to a CDC real-time PCR assay) targeting the tp0967gene of T. pallidum has also been reported [31].
A TPHD-RPA assay, which is based on the RPA technology, was developed by Frimpong et al. [52] to simultaneously and rapidly detect H. ducreyi and T. pallidum. The genes targeted were pol A for yaws and the hemolytic cytotoxin HhdA gene for H. ducreyi. The assay was demonstrated to have 94–95% and 100% sensitivity and specificity, respectively [52] (Frimpong et al., 2020). RPA technology appears to have some advantage over LAMP in that it has a shorter turn-around time (15 min as compared to 30 to 60 min in the case of LAMP) at 37 to 42 degrees [53][54]. To some extent, the technology has been successfully utilized outside typical lab settings in low-resource environments [55][56][57].

3. Molecular Techniques Applicable for the Diagnosis of Buruli Ulcer

Buruli ulcer disease (BUD), caused by Mycobacterium ulcerans (MU), is a skin infection that results in leg and arm ulcers and, if left untreated, can permanently disfigure affected individuals. The disease is generally found in the tropics as well as subtropics, such as west Africa and Asia [58][59]. Transmission is more frequent amongst people residing close to water bodies [60]. Currently, the main laboratory methods used in diagnosing/investigating BUD are culture, microscopy, histopathology, and nucleic acid detection methods such as PCR [15][16][17][61][62]. To confirm BUD diagnosis, the WHO recommends two laboratory tests or one positive microscopy/PCR test in endemic areas [63] (WHO, 2008a; WHO, 2008b).
Despite existence of various diagnostic methods for BUD, PCR is the accepted gold standard. Samples that can be used for PCR include swabs, fine-needle aspirates, and tissue specimens [64][65][66]. The PCR technique specifically targets the sequence referred to as IS2404, giving it its high sensitivity and specificity [67]. Accordingly, the WHO indicates that a positive PCR test result is regarded as enough evidence to start an anti-mycobacterial regimen [63] (WHO, 2008a). It might, however, not be the best tool (compared to culture) to monitor treatment success, as it has been found that the presence of MU DNA persists long after lesions have been treated [68].
Conventional PCR, nested PCR, and real-time PCR have been used for BUD investigation, targeting a number of sequences, including IS2404, IS2606, hsp65, rpoB gene, and 16srRNA gene [60][61][69][70][71][72][73][74][75][76]. Some of the genes targeted are genus- rather than species-specific, and therefore, this necessitates the need to combine with other methods such as restriction fragment length polymorphism (RFLP), sequencing, and oligospecific capture plate hybridization for species differentiation [74][75][76]. Nevertheless, PCR targeting IS2404 has been shown to be more specific, with real-time PCR being more sensitive as compared to conventional PCR [60][66][70][71][77]. Moreover, real-time PCR reduces the possibility of contamination with amplicons from previous reactions. Other techniques employed for BU investigation include LAMP assays (targeting the IS2404 sequence among others) [67][78][79][80], with sensitivities comparable to general conventional PCR but not to real-time PCR [79][80]. A major challenge surrounding the use of LAMP is its adaptability for use in field settings with respect to generating isothermal conditions as well as performing nucleic acid extraction and purification. Thus, the development of DRB-LAMP that utilizes lyophilized reagents and with sensitivity comparable to that of conventional LAMP assay is a laudable approach [81]
A recently developed assay, the RPA for BU diagnosis (targeting the IS2404 sequence) [82], which has short turn-around time. operates at lower isothermal temperatures (compared to LAMP), and has appreciably high specificity and sensitivity of 100% and 88%, respectively, compared to real-time PCR, represents a significant achievement with regards to BU molecular diagnosis.

4. Molecular Techniques Applicable for the Diagnosis of Human African Trypanosomiasis (HAT)/Sleeping Sickness

Sleeping sickness is a parasitic illness mainly spread by tsetse flies. It is caused by two protozoan parasites from the Trypanosoma genus, resulting in two forms of the disease—Trypanosoma brucei gambiense and Trypanosoma brucei rhodesiense [83]. Trypanosoma brucei (T.b.) gambiense is mainly found in Western and Central Africa, accounting for a majority of cases and can be prolonged for months or years [84]. Trypanosoma brucei (T.b.) rhodesiense is mainly found in Eastern and Southern Africa, where it accounts for a minority of cases, causes acute infections, and is considered zoonotic [84][85][86]. Clinical features of the disease can mimic that of other diseases such as malaria. As such, laboratory testing of any suspected case is imperative [87]. In general, diagnoses of HAT is divided into three steps, namely screening, followed by confirmation, and then staging [83]. For screening and confirmation, an antibody-based card agglutination test (CATT/T. b. gambiense), microscopy, as well as RDTs have been used [11][88][89]. However, issues with sensitivities and specificities necessitate the need to also include molecular methods among assays used to diagnose the disease, and a number of these that either detect DNA or RNA have been developed.
Conventionally, nested and real-time PCR assays have been employed that target sequences such as ITS1 DNA and ESAG6/7 gene satellite DNA, among others [90][91][92]. These targets are generally not sub-species-specific. A few sub-species PCR assays in use target sequences such as the TgsGP and SRA gene [93][94][95][96][97][98]. However, these specific assays are generally less sensitive in that the target sequences have relatively fewer copy numbers [93][94][95]. Other assays for subspecies differentiation targeting the SRA and TgsGP sequences have also been used and shown to be more sensitive than their PCR counterparts, which is very encouraging [99][100][101][102].

This entry is adapted from the peer-reviewed paper 10.3390/cimb44100300

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