Fluorescence In Situ Hybridization (FISH) Tests for Identifying Protozoan and Bacterial Pathogens in Infectious Diseases

Diagnosing and treating many infectious diseases depends on correctly identifying the causative pathogen. Characterization of pathogen-specific nucleic acid sequences by PCR is the most sensitive and specific method available for this purpose, although it is restricted to laboratories that have the necessary infrastructure and finance. Microscopy, rapid immunochromatographic tests for antigens, and immunoassays for detecting pathogen-specific antibodies are alternative and useful diagnostic methods with different advantages and disadvantages. Detection of ribosomal RNA molecules in the cytoplasm of bacterial and protozoan pathogens by fluorescence in-situ hybridization (FISH) using sequence-specific fluorescently labelled DNA probes, is cheaper than PCR and requires minimal equipment and infrastructure. A LED light source attached to most laboratory light microscopes can be used in place of a fluorescence microscope with a UV lamp for FISH. A FISH test hybridization can be completed in 30 min at 37 °C and the whole test in less than two hours. FISH tests can therefore be rapidly performed in both well-equipped and poorly-resourced laboratories. Highly sensitive and specific FISH tests for identifying many bacterial and protozoan pathogens that cause disease in humans, livestock and pets are reviewed, with particular reference to parasites causing malaria and babesiosis, and mycobacteria responsible for tuberculosis.


Background
The in-situ hybridization (ISH) technique for examining the formation and detection of RNA-DNA or DNA-DNA nucleotide complementary hybrids in cells utilizing radioactively labelled oligonucleotides as probes was first described in 1969 [1,2]. ISH permits the detection of nucleic acids in individual cells that contain specific nucleotide sequences among a heterogenous population of cells. It also allows the simultaneous determination of biochemical and morphological characteristics of the reactive cells. The hybridization of fluorescently labeled, chromosome-specific, composite DNA probe pools to cytological preparations, termed chromosome painting, has made major contributions to karyotyping and identifying chromosomal changes responsible for human pathology [3]. Fluorescence in situ hybridization (FISH) methods have since been developed to study chromosomal genomic changes at the kilobase level [4]. ISH was first utilized for bacteriology in 1983 with radioactively labeled DNA probes targeting ribosomal RNA (rRNA) [5]. Fluorescently labeled probes have subsequently replaced radioactive probes for FISH [6][7][8]. In 1989, DeLong and colleagues demonstrated that oligodeoxynucleotide probes, complementary to 16S rRNA, labelled with different fluorescent molecules used in FISH can detect single microbial cells and differentiate closely related organisms [9,10]. Shah et al. in 1990 established that FISH can detect and differentiate Pneumocystis carinii strains in sputum and tissue from patients [11]. FISH assays for detecting pathogens in clinical samples now use either peptide nucleic acid (PNA) probes (in which the sugar phosphate backbone is replaced with a more hydrolysis-resistant polyamide chain), locked nucleic acid (LNA) probes (where greater stability is achieved by a methylene bridge linking the 2 oxygen to the 4 carbon of the pentose) or, more commonly, DNA probes [12][13][14][15][16][17][18][19][20].
The application of FISH assays for detecting and identifying microbial pathogens has advanced considerably since the turn of the century [21]. FISH techniques have been applied to investigate the localization of viral nucleic acids within infected tissues and organs, e.g., for SARS-CoV-2 [22,23] and HIV [24], and rarely for identifying the infecting virus [25]. PCR tests, and the detection of viral antigens and specific antibodies to viral antigens, are more commonly and effectively used for diagnosing viral infections. In contrast, FISH tests have proved useful for identifying bacterial, fungal and protozoan disease-causing pathogens, particularly at the species level. Recent examples are listed in Table 1.

Pathogen Type
Test Targets References
Ribosomal RNA molecules possess genus, species and strain-specific regions. Hence, 16S rRNA sequences have been used to establish phylogenetic relationships among bacteria [47]. The binding of rRNA-targeting probes can be visualized without nucleic acid-based Diagnostics 2022, 12, 1286 3 of 18 amplification (NAA) of the target rRNA sequence because rRNA is present in each of the numerous ribosomes in the cytoplasm. Many FISH tests listed in Table 1 are based on DNA or PNA probes that hybridize to specific rRNA sequences in suitably permeabilized cells. Due to the complex three-dimensional structure of rRNA, not all nucleotide sequences within an rRNA molecule are equally accessible for hybridizing with FISH probes. Loop and hairpin formation as well as rRNA-protein interactions hinder hybridization and produce differential binding sensitivity with oligonucleotide probes [48,49]. It is therefore necessary to evaluate and optimize every newly designed probe with the respective reference organism and appropriate negative controls, before it is applied to test samples. Self-annealing and hairpin formation occurring within an oligonucleotide probe itself can lead to low signal intensities, and hence newly designed oligonucleotides also need to be checked for internal complementarity using appropriate software.
The World Health Organization estimated 241 million cases of malaria and 627,000 resulting deaths worldwide in 2020 [50]. Infection with Mycobacterium tuberculosis (MTB), which is primarily responsible for human tuberculosis, caused an estimated 1.4 million deaths worldwide in 2020 [51]. Malaria and tuberculosis can manifest as latent infections that rapidly become fulminant diseases with fatal consequences. Babesiosis is a potentially fatal, tick-borne, globally emerging human disease, that also afflicts livestock and domestic pets [52]. While the principle of FISH tests remains the same, their application to identify different pathogens can vary considerably. Recently developed FISH tests that can be easily used in resource-limited laboratories worldwide for identifying causative pathogens in malaria, tuberculosis and babesiosis are therefore selected for detailed consideration in this article. These FISH tests for malaria, tuberculosis and babesiosis have the following shared characteristics: (i) the assays are performed on thin smears on glass microscope slides, (ii) cells in the smear are rapidly permeabilized for hybridization which is then performed with fluorescently labelled DNA probes at 37 • C for 30 min, (iii) fluorescence can be viewed under LED illumination in common light microscopes as shown in Supplementary Figure S1, (iv) the test is completed in less than two hours, and (v) only living cells are labelled because rRNA is rapidly degraded in dying cells. Supplementary Figure S2 summarizes the published work-flow for tuberculosis FISH tests [28,29].

Background
Plasmodium falciparum is responsible for most of the annual 241 million global malaria infections, together with an estimated 4.5 million cases of Plasmodium vivax and fewer cases of Plasmodium malariae and Plasmodium ovale [50]. Plasmodium knowlesi, which normally infects macaque and leaf monkeys, causes a significant number of dead-end human infections in Southeast Asian countries [53]. Plasmodium knowlesi is difficult to differentiate from human malaria parasites in Giemsa-stained blood smears that are commonly used for diagnosing malaria in endemic areas [53]. Occasional zoonotic infections with Plasmodium cynomolgi and Plasmodium inui have also been reported in Southeast Asia [53]. Plasmodium brasilianum and Plasmodium simium infect platyrrhine monkeys in South and Central America, are genetically almost identical to the human malaria parasites P. malariae and P. vivax respectively, and likely to have been derived from the human parasites by anthroponosis [53]. P. simium and P. brasilianum can also infect humans by zoonosis but their differentiation from P. vivax and P. malariae respectively in stained blood smears is not possible [53].
There were 2171 US cases of malaria reported to the US Centers for Disease Control and Prevention (CDC) in 2017 according to the latest available CDC report [54]. Plasmodium falciparum accounted for 70.5%, P. vivax 10.0%, P. ovale 5.5%, and P. malariae 2.6% of the infections, all of which had been acquired outside the US [54]. Infections with two or more Plasmodium species were responsible for 1.0% of infections [54]. The identification of malaria parasites for diagnosis is therefore also needed in the US and other countries with no indigenous transmission of malaria.
Giemsa-stained thick and thin blood smear microscopy has been the most widely used technique globally for diagnosing malaria. It however requires time and an experienced microscopist for optimal sensitivity of detection and for identifying the infecting species of parasite. A sensitivity of >150 parasites per µL is typically achieved during routine microscopy [39,55]. Rapid diagnostic tests (RDTs), based on the immunochromatographic detection of antibodies to the histidine rich protein 2 of P. falciparum (PfHRP2) and pan Plasmodium-specific lactate dehydrogenase and aldolase, have more recently proved helpful in resource-limited locations [55]. However, selection for PfHRP2 gene deletions in P. falciparum in malaria-endemic areas of Africa has lately increased the false negativity rates for P. falciparum [55]. PCR-dependent NAA diagnostic tests have the best sensitivity and specificity and are able to identify Plasmodium at the species level, but the procedure is not suitable for resource-poor settings and field use [56]. The Loop-Mediated Isothermal Amplification (LAMP) technique has the desired sensitivity and specificity but is not widely utilized for routine malaria diagnosis and species identification [57]. Flow cytometric detection of malaria parasites in blood have also been recently described although details of its limit of detection and ability to identify different species remain to be established [58,59].

Genus-Specific FISH Test That Identifies All Common Human Malaria Parasites
Simple, rapid and specific FISH tests for malaria, that can be easily performed in resource-constrained diagnostic laboratories, have many advantages [39,42]. These FISH tests for malaria employ a similar protocol to FISH tests for tuberculosis shown in Figure S2. A standard laboratory microscope with a LED fluorescence unit attached to it can be used to read the processed smears on the slide ( Figure S1). Results from a Plasmodium genus-specific FISH test utilizing a DNA probe hybridizing to 18S rRNA [39] are reproduced in Figure 1.  The Plasmodium genus-specific FISH test identified all common species of human malaria parasites with 100% specificity when compared with several other common human blood-borne pathogens [39].

FISH Test for Specifically Identifying Plasmodium Falciparum
A PF-FISH test that complements the Plasmodium genus-specific FISH test, and designed to specifically identify P. falciparum, utilized a mixture of P. falciparum 18S rRNA-specific probes labeled with Alexa 488 green and the Plasmodium genus-specific probe labeled with a Texas Red in a multiplex format. Plasmodium falciparum fluoresced green ( Figure 2), while all Plasmodium parasites, including P. falciparum, fluoresced red with appropriate light filters in the PF-FISH test [39].

FISH Test for Specifically Identifying Plasmodium Vivax
In a second complementary FISH test, termed the PV-FISH test, a mixture of DN probes that hybridize only to the 18S rRNA of P. vivax were labeled with the Alexa 48

FISH Test for Specifically Identifying Plasmodium Vivax
In a second complementary FISH test, termed the PV-FISH test, a mixture of DNA probes that hybridize only to the 18S rRNA of P. vivax were labeled with the Alexa 488 green, and used in a multiplex format with the Plasmodium genus-specific probe labeled with Texas Red. Only P. vivax fluoresced green, and all Plasmodium species fluoresced red, with appropriate filters in the PV-FISH test [39] as shown in Figure 3.
The two FISH tests for specifically identifying P. falciparum and P. vivax had greater analytical sensitivity, and also higher clinical sensitivity and specificity, compared to microscopic examination of Giemsa-stained blood smears [39]. green, and used in a multiplex format with the Plasmodium genus-specific probe labe with Texas Red. Only P. vivax fluoresced green, and all Plasmodium species fluoresced r with appropriate filters in the PV-FISH test [39] as shown in Figure 3. The two FISH tests for specifically identifying P. falciparum and P. vivax had grea analytical sensitivity, and also higher clinical sensitivity and specificity, compared to croscopic examination of Giemsa-stained blood smears [39].

FISH Test for Specifically Identifying Plasmodium Knowlesi
Zoonotic P. knowlesi infections in Southeast Asia are commonly misidentified a malariae or P. falciparum in Giemsa-stained human blood smears because of morpholog similarities between the blood stages, so that PCR-based tests were needed for confirm P. knowlesi infections [60][61][62][63][64][65]. Correct diagnosis of P. knowlesi malaria is essential for t reasons: (i) understanding its epidemiology, and (ii) its pathogenicity and drug treatm options can differ from human malaria caused by P. falciparum, P. malariae, P. ovale and vivax that also occur in Southeast Asia. Reliable RDTs for specifically detecting P. know are not yet available [66]. However, a simple and specific FISH test using DNA pro targeting P. knowlesi 18S rRNA (termed the PK-FISH test) specifically identified P. know in blood smears [42], as shown in Figure 4.

FISH Test for Specifically Identifying Plasmodium Knowlesi
Zoonotic P. knowlesi infections in Southeast Asia are commonly misidentified as P. malariae or P. falciparum in Giemsa-stained human blood smears because of morphological similarities between the blood stages, so that PCR-based tests were needed for confirming P. knowlesi infections [60][61][62][63][64][65]. Correct diagnosis of P. knowlesi malaria is essential for two reasons: (i) understanding its epidemiology, and (ii) its pathogenicity and drug treatment options can differ from human malaria caused by P. falciparum, P. malariae, P. ovale and P. vivax that also occur in Southeast Asia. Reliable RDTs for specifically detecting P. knowlesi are not yet available [66]. However, a simple and specific FISH test using DNA probes targeting P. knowlesi 18S rRNA (termed the PK-FISH test) specifically identified P. knowlesi in blood smears [42], as shown in Figure 4.  The PK-FISH test, like the analogous Plasmodium genus-specific test and the PF-FI and PV-FISH tests [39], identified all asexual blood stages, i.e., rings, trophozoites a schizonts, as shown in Figure 5 for P. knowlesi [42]. The PK-FISH test also detected knowlesi at the low limit of 16 P. knowlesi parasites per µL, even in the concomitant p . Specificity of the PK-FISH test for P. knowlesi. Photographs showing PK-FISH test results with the P. knowlesi-specific probe (green fluorescence) and the Plasmodium genus-specific probe (orange fluorescence) in blood smears containing P. knowlesi from monkey blood (Pk), and from human blood with confirmed infections of P. falciparum (Pf), P. malariae (Pm), P. ovale (Po) and P. vivax (Pv).
Each set of paired photographs shows fluorescence in the same field when viewed in a LED fluorescence microscope with appropriate light filters ( Figure S1). The scale bars represent approximately 5 µm. Reproduced with permission under the creative commons license from Reference [42].
The PK-FISH test, like the analogous Plasmodium genus-specific test and the PF-FISH and PV-FISH tests [39], identified all asexual blood stages, i.e., rings, trophozoites and schizonts, as shown in Figure 5 for P. knowlesi [42]. The PK-FISH test also detected P. knowlesi at the low limit of 16 P. knowlesi parasites per µL, even in the concomitant presence of P. falciparum at approximately 500 parasites per µL [42], which is superior to that possible with routine microscopic examination of Giemsa-stained thin blood films [39,55]. This property is very useful in Southeast Asia where mixed infections of P. knowlesi and other human malaria parasite species are common [60,63]. The highly specific and sensitive PK-FISH therefore meets a widely-recognized diagnostic need of peripheral and district-level clinical laboratories in areas of Southeast Asia where P. knowlesi zoonosis is prevalent [53,[60][61][62][63][64][65][66].
(orange fluorescence) in blood smears containing P. knowlesi from monkey blood (Pk), and from human blood with confirmed infections of P. falciparum (Pf), P. malariae (Pm), P. ovale (Po) and P vivax (Pv). Each set of paired photographs shows fluorescence in the same field when viewed in LED fluorescence microscope with appropriate light filters ( Figure S1). The scale bars represent ap proximately 5 µm. Reproduced with permission under the creative commons license from referenc 42.
The PK-FISH test, like the analogous Plasmodium genus-specific test and the PF-FISH and PV-FISH tests [39], identified all asexual blood stages, i.e., rings, trophozoites and schizonts, as shown in Figure 5 for P. knowlesi [42]. The PK-FISH test also detected P knowlesi at the low limit of 16 P. knowlesi parasites per µL, even in the concomitant pres ence of P. falciparum at approximately 500 parasites per µL [42], which is superior to tha possible with routine microscopic examination of Giemsa-stained thin blood films [39,55 This property is very useful in Southeast Asia where mixed infections of P. knowlesi and other human malaria parasite species are common [60,63]. The highly specific and sensi tive PK-FISH therefore meets a widely-recognized diagnostic need of peripheral and district-level clinical laboratories in areas of Southeast Asia where P. knowlesi zoonosis i prevalent [53,[60][61][62][63][64][65][66]. Photographs showing results from the PK-FISH test with the P. knowlesi-specific probe (green fluorescence) and the Plasmodium genus-specific probe (orange fluorescence) on R-rings; T-trophozoites; S-schizonts. Dual colour fluorescence in the same field is shown in paired photographs R1 and R2, T1 and T2, and S1 and S2. Fluorescence was viewed in a LED fluorescence microscope with pertinent light filters ( Figure S1). The ring, trophozoite and schizont-stage parasites were produced from synchronised in vitro cultures of P. knowlesi. Parasites stained with Giemsa from smears prepared in parallel to the corresponding smears used in the PK-FISH test are shown in R3, T3 and S3 respectively. The scale bars represent approximately 5 µm. Reproduced with permission under the creative commons license from Reference [42].

Conclusions
A large number of malaria tests are performed for diagnostic and screening purposes in malaria-endemic countries. Tests in malaria-free countries are utilized for (i) screening passengers arriving from malaria-endemic countries to prevent the reintroduction of malaria if mosquito vectors are present in the country of arrival, and (ii) confirming malaria in arriving travelers who have malaria-like symptoms. FISH tests are more costly and complex to perform than Giemsa-stained blood smear microscopy and RDTs, but significantly less so than NAA-dependent PCR and LAMP, for detecting Plasmodium infections (table in Section 5 below). FISH tests are particularly useful for identifying the species of infecting parasites, as illustrated here for P. falciparum, P. vivax and P. knowlesi. Therefore, the clinical diagnostic characteristics and simple methodology of the newly described FISH tests for malaria parasites, suggest that they can (i) usefully complement Giemsa-stained blood smear microscopy and RDTs for routine diagnosis and screening for malaria, and (ii) identify the species of infecting Plasmodium (including in mixed infections), in both endemic and non-endemic countries.
Laboratory tests commonly used for diagnosing babesiosis involve the detection of (i) parasites in stained blood smears by microscopy, (ii) serum antibodies to Babesia by immunoassays, and (iii) Babesia-specific nucleic acid sequences by PCR [69,70]. Babesiosis and borreliosis (a tick vector-borne disease caused by spirochete Borrelia bacteria), share many clinical manifestations, and occur as coinfections [72][73][74][75][76]. They have an overlapping geographical distribution [67][68][69][70][71]77,78], underscoring the importance of diagnostic laboratory tests for differentiating babesiosis and borreliosis. Early intra-erythrocytic stages of human-infecting Babesia species are not readily distinguished from the ring and trophozoite stages of P. falciparum by microscopy in areas where babesiosis and malaria are co-endemic [69]. Furthermore, the antibody assays for diagnosing human and veterinary babesiosis cannot easily differentiate between active and resolved Babesia infections [67][68][69][70]. PCR tests for babesiosis have high sensitivity [79][80][81] and are recommended for screening donor blood for babesiosis in the US [82]. However, cost and infrastructure requirements make PCR-based tests impractical for use in resource-limited laboratories and field settings.

Babesia Genus-Specific FISH Test
A FISH test that identifies all common species of Babesia parasites, with many advantages for use in resource-limited laboratories, has been developed [43,44]. Termed the Babesia genus FISH test, it is based on DNA probes that specifically hybridize to the multiple copies of Babesia 18S rRNA present in the parasite cytoplasm. Like other rRNA-directed FISH tests, the Babesia genus FISH test does not require NAA-a process that is sensitive to NAA inhibitors sometimes present in blood [83].
The Babesia genus-specific FISH test detects B. microti, B. duncani and B. divergens, as well as the two important parasites causing bovine babesiosis, B. bovis and B. bigemina [43], as illustrated in Figure 6.
The Babesia genus-specific FISH test, in conjunction with an IFA test for detecting serum antibodies to B. duncani and B. microti, on clinical samples originating from USA, Australia, Europe and elsewhere, showed that the global prevalence of B. duncani infections had hitherto been under-estimated [44]. Furthermore, the Babesia genus-specific FISH test was highly specific and did not detect other pertinent pathogens found in human blood [43], including different species of Borrelia and Plasmodium [43], as well as various species of Bartonella that infect humans and domestic pets [33,84].
The Babesia genus-specific FISH test detects B. microti, B. duncani and B. diverg well as the two important parasites causing bovine babesiosis, B. bovis and B. bigemi as illustrated in Figure 6.
The Babesia genus-specific FISH test, in conjunction with an IFA test for de serum antibodies to B. duncani and B. microti, on clinical samples originating from Australia, Europe and elsewhere, showed that the global prevalence of B. duncan tions had hitherto been under-estimated [44]. Furthermore, the Babesia genus-s FISH test was highly specific and did not detect other pertinent pathogens found man blood [43], including different species of Borrelia and Plasmodium [43], as well ious species of Bartonella that infect humans and domestic pets [33,84].

Conclusions
The prevalence of human babesiosis has probably been underestimated throu the world [44]. Babesiosis also afflicts livestock and pets [52]. The clinical diagnosti acteristics and simple methodology of the FISH tests show that they can complem isting diagnostic methods to meet an increasing need to specifically and easily id Babesia infections in patients and animals. FISH tests can be used in mixed infection is also useful for histopathological investigations to identify Babesia parasites seque in tissues [45]. Species-specific Babesia FISH tests, that are presently being develop address more precise diagnostic requirements in babesiosis.

Conclusions
The prevalence of human babesiosis has probably been underestimated throughout the world [44]. Babesiosis also afflicts livestock and pets [52]. The clinical diagnostic characteristics and simple methodology of the FISH tests show that they can complement existing diagnostic methods to meet an increasing need to specifically and easily identify Babesia infections in patients and animals. FISH tests can be used in mixed infections. FISH is also useful for histopathological investigations to identify Babesia parasites sequestered in tissues [45]. Species-specific Babesia FISH tests, that are presently being developed, can address more precise diagnostic requirements in babesiosis.

Background
Pulmonary mycobacterial infections in humans are caused mostly by Mycobacterium tuberculosis (MTB) and the closely related species Mycobacterium bovis, both of which belong to the Mycobacterium tuberculosis complex (MTBC) [51]. Infections with non-tuberculous mycobacteria (NTM), including the Mycobacterium avium complex (MAC), M. kansasii, M. fortuitum, M. xenopi, M. abscessus, and M. simiae also occur worldwide, making their differential diagnosis important for clinical purposes [51,[85][86][87]. Infections with MAC are common in late-stage human immunodeficiency virus infections, where the mycobacteria are often restricted to lymphoid tissue. FISH provides a sensitive and specific method for detecting MAC by in biopsied tissues, which is important because the management and treatment of patients with MTBC and NTM infections are different [30]. Norcardiosis, caused by related Norcadia species widely distributed in the environment, also needs to be differentiated from MTB during lung infections [88].
Microscopic examination for acid-fast staining (AFS) bacilli, e.g., with the Ziehl-Neelsen stain, in sputum or tissue plays an important role in the diagnosis of tuberculosis [89]. AFS does not differentiate between mycobacterial species. It also lacks sufficient sensitivity with sputum smears and tissue samples. Sensitivity is increased in sputum smears and biopsied tissue by staining with auramine and detecting fluorescence in a LED fluorescence microscope [90], similar to that used for FISH tests ( Figure S1). The Xpert ® MTB/RIF system or Xpert (Cepheid, Sunnyvale, CA, USA), is a PCR-based nucleic acid amplification (NAA) technique that detects specific DNA sequences of MTB [91][92][93]. Xpert is recommended by the WHO for identifying MTB and rifampicin resistance in the sputum of adults and children presumed to have tuberculosis [91][92][93]. It is approximately 100 times more sensitive for detecting MTB than conventional AFS, but Xpert identifies only MTB, and its use in many resource-constrained endemic countries is limited by cost. Culturing clinical specimens continues to have an important role in identifying infecting mycobacteria in tuberculosis-like disease, especially in smear negative, pediatric or extra pulmonary infections and resource-limited laboratories. Culture techniques that significantly reduce culture times for identifying mycobacteria are becoming available to facilitate diagnosis [94]. Immunochromatographic tests to detect specific proteins produced by MTBC are more cost-effective than PCR tests, but, as yet, do not have the desired specificity and sensitivity [92,93].

FISH Tests for Identifying the Genus Mycobacterium as Well as the Mycobacterium Tuberculosis and Mycobacterium Avium Complexes in Culture
A simple and rapid test for directly identifying MTB and NTM in sputum and tissues that can be used by resource-limited laboratories in endemic countries is therefore expected to greatly aid tuberculosis control worldwide [92,93]. Two dual color FISH tests, with simple protocols (Figure S2), and requiring only a LED fluorescence microscope ( Figure S1), meet this need [28][29][30]. The MN Genus-MTBC FISH test used an orange fluorescent DNA probe that specifically hybridizes to the 23S rRNA of the Mycobacterium tuberculosis complex (MTBC) and a green fluorescent probe specific for the Mycobacterium and Nocardia genera (MN Genus) 16S rRNA to detect and distinguish MTBC from other mycobacteria and Nocardia species. A complementary MTBC-MAC FISH test used green and orange fluorescent probes for 23S rRNA that respectively differentiate MTBC and MAC [28][29][30].
All Mycobacterium species from reference cultures, except M. wolinskyi, reacted positively with the MN Genus-specific probe and only the M. tuberculosis complex species reacted positively with the MTBC-specific probe in the MN Genus-MTBC FISH test. Only the M. tuberculosis complex species reacted positively with the MTBC-specific probe and only the M. avium complex species reacted positively with the MAC-specific probe in the MTBC-MAC FISH test [28]. Nocardia reacted positively with the MN Genus probe but not with the MTBC-and MAC-specific probes in the MN Genus-MTBC and the MTBC-MAC tests [28]. The estimated specificity of the two FISH tests for MTBC and MAC in reference cultures was 100%, with a limit of detection of 1.5-5.1 × 10 4 bacteria per ml [28]. Results from the two FISH tests with reference strain cultures of M. tuberculosis, M. avium and M. kansasii [28] are reproduced in Figure 7

FISH Tests for Identifying MTBC and MAC in Sputum
The FISH tests used for culture identification can also be used for directly detecting mycobacteria in sputum [29]. with the MTBC-and MAC-specific probes in the MN Genus-MTBC and the MTBC tests [28]. The estimated specificity of the two FISH tests for MTBC and MAC in ref cultures was 100%, with a limit of detection of 1.5-5.1 × 10 4 bacteria per ml [28]. R from the two FISH tests with reference strain cultures of M. tuberculosis, M. avium a kansasii [28] are reproduced in Figure 7.

FISH Tests for Identifying MTBC and MAC in Sputum
The FISH tests used for culture identification can also be used for directly det mycobacteria in sputum [29]. Figure 8 reproduces results obtained with the MN G MTBC FISH test performed directly on a sputum smear containing MTBC which r with the MN Genus-and MTBC-specific probes and a different smear containing scessus, an NTM, that reacted only with the MN Genus-specific probe.

Other FISH Tests for Tuberculosis
Another FISH test specific for MTBC in sputum targeting the rpoB gene codi the β subunit of RNA polymerase has been described [95]. The rpoB FISH test ho required enzyme digestion of sputum, concentration of mycobacteria by centrifug hybridization overnight and a UV fluorescence microscope for visualizing result Other PNA or DNA probe-based FISH tests described for MTBC and MAC [12-16 require long and more stringent hybridization procedures, and a UV fluorescence scope for viewing test results. They have not been used yet for routine diagnosis demic countries. The MN Genus-MTBC and MTBC-MAC FISH tests on the other cost < US$5 per test, provide results in <2 h after sputum collection, do not require en treatment and centrifugation, can use LED fluorescence microscopy ( Figure S1), an lize reagents that are stable at ambient temperature [29]. They are used in India [3 FISH assays have the advantage that they are unaffected by inhibitors in respiratory ples which reduce sensitivity and require elaborate controls for NAA tests [96,97].

Conclusions
FISH tests for tuberculosis meet internationally-expressed needs for diagnosing tuberculosis in respiratory samples [92,93]. They can complement AFS microscopy and NAA methods to detect and differentiate MTBC from MAC and other NTM in sputum

Other FISH Tests for Tuberculosis
Another FISH test specific for MTBC in sputum targeting the rpoB gene coding for the β subunit of RNA polymerase has been described [95]. The rpoB FISH test however required enzyme digestion of sputum, concentration of mycobacteria by centrifugation, hybridization overnight and a UV fluorescence microscope for visualizing results [95]. Other PNA or DNA probe-based FISH tests described for MTBC and MAC [12][13][14][15][16] also require long and more stringent hybridization procedures, and a UV fluorescence microscope for viewing test results. They have not been used yet for routine diagnosis in endemic countries. The MN Genus-MTBC and MTBC-MAC FISH tests on the other hand, cost < US$5 per test, provide results in <2 h after sputum collection, do not require enzyme treatment and centrifugation, can use LED fluorescence microscopy ( Figure S1), and utilize reagents that are stable at ambient temperature [29]. They are used in India [30]. All FISH assays have the advantage that they are unaffected by inhibitors in respiratory samples which reduce sensitivity and require elaborate controls for NAA tests [96,97].

Conclusions
FISH tests for tuberculosis meet internationally-expressed needs for diagnosing tuberculosis in respiratory samples [92,93]. They can complement AFS microscopy and NAA methods to detect and differentiate MTBC from MAC and other NTM in sputum and cultures. FISH is also useful in detecting MTB and MAC in biopsied tissues [30]. Table in Section 5 below compares NAA and FISH tests for tuberculosis.

Comparison of NAA and FISH Tests for Diagnosing Malaria and Tuberculosis
Tests that depend on amplifying specific nucleic acid sequences of pathogens and the subsequent detection and/or sequencing of the amplified nucleic acids are widely regarded as the gold standard for diagnostic tests because of their high sensitivity and specificity. Two NAA techniques that can be used for RNA and DNA, are based on PCR [98] and LAMP [99]. Diagnostic test needs vary considerably for different pathogens and the diseases caused by them. Malaria and tuberculosis are parasitic and bacterial diseases respectively of great global clinical concern [50,51]. The use of PCR and LAMP tests for identifying pathogens causing the two diseases are therefore compared with FISH tests in Table 2. Light microscope with LED/filter attachment ( Figure S1); 37 • incubator. Low maintenance cost.

Personnel
Highly trained operator for PCR and LAMP. Trained microscopist. Individual samples and not presently automated. Amenable to automation by flow cytometry [24] and fluorescence detection by digital imaging.

Laboratory and Location Suitability
(i) Malaria: PCR rarely used for primary diagnosis except zoonotic malaria. LAMP rarely used for primary diagnosis of malaria.
(ii) Tuberculosis: LAMP comparable to Xpert for tuberculosis [101]. Xpert not advantageous in locations with low levels of multi drug resistant (MDR) M. tuberculosis [101][102][103] or low disease prevalence [103]. LAMP not useful in areas with high levels of MDR [101].
All types of laboratories, locations and field use. Does not presently detect MDR M. tuberculosis.
(ii) Tuberculosis: Xpert only identifies MTB as do common LAMP tests.

Sensitivity to Inhibitors in Clinical Samples
PCR and LAMP sensitive to inhibitors in some tissue and sputum samples [83,96,97]. No FISH inhibitors in clinical samples.

Detection of Live vs. Dead Pathogens
PCR and LAMP detect DNA in both dead and live cells because of DNA stability [109]. Cell morphology remains unknown.
Detects live organisms only because rRNA degrades rapidly in dying cells [39,110]. Cell morphology visible. Useful for monitoring drug treatment & disease course

Overall Conclusions and Future Prospects
The simplicity, cost, modest infrastructure/equipment/reagent requirements, reagent stability, good diagnostic parameters, and the ability to identify pathogens at the species level, suggest that FISH tests can be used in advanced as well as resource-constrained diagnostic laboratories throughout the world. FISH tests, therefore, can complement existing diagnostic tests in both disease-endemic and non-endemic countries. FISH tests are particularly advantageous for identifying pathogens at the species level. Flow cytometry combined with FISH has been shown able to rapidly identify potentially pathogenic bacteria present in food, water, air and biofilms formed on various abiotic surfaces [110]. The use of the Flow-FISH methodology [24,110] for identifying causative pathogens in infectious diseases therefore merits further investigation. FISH may also be usefully interfaced with advanced optical and microscopic techniques [22,23,25,111,112] to further expand its scope for identifying infecting pathogens for research and diagnostic purposes.
Author Contributions: J.S.S. and R.R. wrote and approved the manuscript. All authors have read and agreed to the published version of the manuscript.