Are nanobiosensors an improved solution for diagnosis of Leishmania?

pharmaceutics Review Are Nanobiosensors an Improved Solution for Diagnosis of Leishmania? Sona Jain 1,* , Wanessa Santana 1, Silvio S. Dolabella 2, AndréL. S. Santos 3, Eliana B. Souto 4,5,* and Patrícia Severino 1   Citation: Jain, S.; Santana, W.; Dolabella, S.S.; Santos, A.L.S.; Souto, E.B.; Severino, P. Are Nanobiosensors an Improved Solution for Diagnosis of Leishmania?. Pharmaceutics 2021,13, 491. https://doi.org/10.3390/ pharmaceutics13040491 Academic Editor: Tomáš Etrych Received: 15 March 2021 Accepted: 1 April 2021 Published: 3 April 2021 Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations. Copyright: © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/). 1Postgraduate Program in Industrial Biotechnology, Universidade Tiradentes, Aracaju 49032-490, Brazil; [email protected] (W.S.); [email protected] (P.S.) 2 Department of Morphology, Federal University of Sergipe, São Cristóvão 49100-000, Brazil; [email protected] 3Paulo de Góes Microbiology Institute, Departament of General Microbiology, Federal University of Rio de Janeiro, Rio de Janeiro 21941-901, Brazil; andr[email protected] 4CEB—Centre of Biological Engineering, University of Minho, Campus de Gualtar, 4710-057 Braga, Portugal 5 Department of Pharmaceutical Technology, Faculty of Pharmacy, University of Coimbra, Pólo das Ciências da Saúde, Azinhaga de Santa Comba, 3004-531 Coimbra, Portugal *Correspondence: [email protected] (S.J.); [email protected] (E.B.S.) Abstract: Leishmaniasis is one of the deadliest neglected tropical diseases affecting 12–15 million people worldwide, especially in middleand low-income countries. Rapid and accurate diagnosis of the disease is important for its adequate management and treatment. Several techniques are available for the diagnosis of leishmaniasis. Among these, parasitological and immunological tests are most widely used. However, in most cases, the utilized diagnostic techniques are not good enough, showing cross-reactivity and reduced accuracy. In recent years, many new methods have been reported with potential for improved diagnosis. This review focuses on the diagnosis of Leishmania exploring the biosensors and nanotechnology-based options for their detection. New developments including the use of nanomaterials as fluorophores, fluorescence quenchers as reducing agents and as dendrimers for signal improvement and amplification, together with the use of aptamers to replace antibodies are described. Future research opportunities to overcome the current limitations on the available diagnostic approaches are also discussed. Keywords: leishmaniasis; nanomaterials; immunosensors; genosensors; parasitic diseases 1. Introduction Leishmaniasis is one of the seven most important tropical diseases in the world. It is caused by the obligate intracellular parasitic protozoa belonging to the genus Leishmania and affects millions of people in the world. In 2018, leishmaniasis was reported in 98 countries and territories across four continents. With approximately 1–2 million new cases and 70,000 deaths every year, it is among the deadliest of the neglected tropical diseases (NTDs) [1,2]. Leishmaniasis usually affects underdeveloped countries in Africa, Asia, and Latin America (Figure 1A), and is related to malnutrition, population migration, poor living conditions, fragile immune system, and lack of resources [ 3 ]. Recent surveys indicate that approximately 350 million people live in a vulnerable situation with the risk of contracting leishmaniasis. From a global perspective, the disease currently affects approximately 12–15 million people worldwide [4]. Pharmaceutics 2021,13, 491. https://doi.org/10.3390/pharmaceutics13040491 https://www.mdpi.com/journal/pharmaceutics Pharmaceutics 2021,13, 491 2 of 19 Pharmaceutics 2021, 13, x FOR PEER REVIEW 2 of 20 edentates, monkeys, and wild or domestic canines (Figure 1B). Humans are accidentally infected in endemic areas [6]. Distinct species of Leishmania can cause different clinical manifestations, and the disease can be grouped into three main clinical forms: cutaneous leishmaniasis (CL), mucocutaneous leishmaniasis (ML), and visceral leishmaniasis (VL), also known as kala-azar [3]. Each form varies in the degree of severity, where VL is the most severe form presenting the highest mortality. Figure 1. Geographical distribution of leishmaniasis in the world (A). Life cycle of Leishmania in sand fly and mammalian host (B). The two main evolutionary forms during their life cycle: the promastigote (in the invertebrate host) and the amastigote (present in the vertebrate host) are shown. CL is the most prevalent form caused by leishmanial species, such as L. tropica, L. amazonensis, L. aethiopica, and L. major, and it is generally painless and chronic, resulting in constant scar and critical stigma or disability [7]. ML is most frequently found in Americas and is caused by species such as L. braziliensis and L. mexicana. ML is characterized by the appearance of lesions that may partially or completely destroy the mucous membranes of the cavities of the nose, mouth, and throat, leading to disfigurement of the patient with consequent social exclusion [8]. As mentioned before, VL is the most severe form of the disease caused by L. donovani and L. infantum, and it is characterized by fever, weight loss, enlargement of liver and spleen, pancytopenia, and hypergammaglobulinemia and may be fatal if left untreated [9]. Early treatment as well as rapid and accurate diagnosis of the disease are important for its control [10,11]. Parasitological, serological, and molecular methods currently in use for diagnosis still have limitations that make the confirmation of the disease difficult [12]. The parasitological diagnosis, in addition to being invasive, shows low sensitivity. Serological tests work well in people who have a good immune response, but immuno-depressed patients, for example, do not respond well due to deficiencies in antibody production. Molecular tests, although of high precision, generally depend on equipment that is not always available [13]. The current diagnostic methods are thus often not sufficient, being presumptive, and diagnosis in most cases usually involves the identification of the Leishmania species mainly based on local epidemiology and clinical aspects [14]. This generates an urgent need to develop more sensitive, specific, and accessible tests, with easy application in the field, especially in poorly or modestly equipped laboratories. Recent years have seen a great advancement in the development of nanomaterials and bioanalytical chemistry. Many emerging methods have been described based on nanomaterials such as optical labels (Hu et al. 2017), use of new signal transduction mechanisms [15], and use of aptamers over antibodies [16,17]. In this review, we discuss the state-ofFigure 1. Geographical distribution of leishmaniasis in the world ( A ). Life cycle of Leishmania in sand fly and mammalian host ( B ). The two main evolutionary forms during their life cycle: the promastigote (in the invertebrate host) and the amastigote (present in the vertebrate host) are shown. There are more than 20 Leishmania species distributed worldwide that are transmitted by over 90 phlebotomine sandfly species [ 5 ]. Leishmaniasis is usually transmitted by the bite of an infected female sandfly to mammalian host, such as rodents, marsupials, edentates, monkeys, and wild or domestic canines (Figure 1B). Humans are accidentally infected in endemic areas [ 6 ]. Distinct species of Leishmania can cause different clinical manifestations, and the disease can be grouped into three main clinical forms: cutaneous leishmaniasis (CL), mucocutaneous leishmaniasis (ML), and visceral leishmaniasis (VL), also known as kala-azar [ 3 ]. Each form varies in the degree of severity, where VL is the most severe form presenting the highest mortality. CL is the most prevalent form caused by leishmanial species, such as L. tropica, L. amazonensis ,L. aethiopica, and L. major, and it is generally painless and chronic, resulting in constant scar and critical stigma or disability [ 7 ]. ML is most frequently found in Americas and is caused by species such as L. braziliensis and L. mexicana. ML is characterized by the appearance of lesions that may partially or completely destroy the mucous membranes of the cavities of the nose, mouth, and throat, leading to disfigurement of the patient with consequent social exclusion [ 8 ]. As mentioned before, VL is the most severe form of the disease caused by L. donovani and L. infantum, and it is characterized by fever, weight loss, enlargement of liver and spleen, pancytopenia, and hypergammaglobulinemia and may be fatal if left untreated [9]. Early treatment as well as rapid and accurate diagnosis of the disease are important for its control [ 10 , 11 ]. Parasitological, serological, and molecular methods currently in use for diagnosis still have limitations that make the confirmation of the disease difficult [ 12 ]. The parasitological diagnosis, in addition to being invasive, shows low sensitivity. Serological tests work well in people who have a good immune response, but immuno-depressed patients, for example, do not respond well due to deficiencies in antibody production. Molecular tests, although of high precision, generally depend on equipment that is not always available [ 13 ]. The current diagnostic methods are thus often not sufficient, being presumptive, and diagnosis in most cases usually involves the identification of the Leishmania species mainly based on local epidemiology and clinical aspects [ 14 ]. This generates an urgent need to develop more sensitive, specific, and accessible tests, with easy application in the field, especially in poorly or modestly equipped laboratories. Recent years have seen a great advancement in the development of nanomaterials and bioanalytical chemistry. Many emerging methods have been described based on nanomaterials such as optical labels (Hu et al. 2017), use of new signal transduction Pharmaceutics 2021,13, 491 3 of 19 mechanisms [ 15 ], and use of aptamers over antibodies [ 16 , 17 ]. In this review, we discuss the state-of-the-art on the use of nanomaterials for the detection of leishmanial parasites and the role of surface-tailored aptamers for the improved diagnosis of the disease. 2. Current Diagnostic Methods and Limitations The control of leishmaniasis requires a combined set of intervention strategies, among which early diagnosis and treatment are important aspects. Current diagnosis is mostly based on the use of immunological and parasitological tests combined with clinical symptoms. However, in the case of CL and ML, serological tests have limited value. 2.1. Parasitological Tests Parasitological tests are still considered the gold standard for the diagnosis of leishmaniasis. In this direct identification method, diagnosis is made microscopically by identifying amastigotes in affected tissues (CL/ML injury site) or in sample aspirated from the spleen, bone marrow, or lymph nodes in case of VL patients [ 10 , 11 , 14 ]. The inoculation of samples in experimental animals (xenodiagnoses), as well as the in vitro culture of Leishmania promastigotes in culture medium, are also used in the routine diagnosis of leishmaniasis. A combination of microscopy and culture methods enhances the diagnostic sensitivity by more than 85% [ 14 , 18 ]. However, all these methods are expensive and time-consuming, require skilled labor, and do not discriminate between Leishmania species [10,11]. 2.2. Immunological Tests The leishmania skin test or Montenegro test is used to measure the delayed-type hypersensitivity reaction to an intradermal injection of a suspension of killed Leishmania promastigotes [ 10 , 11 ]. It is a simple, sensitive, and specific method for CL; however, it does not allow the identification of species and does not differentiate past infections from present infections [ 10 ]. In addition, the test gives negative results during the disease period in VL [11]. Serological tests like indirect immunofluorescence assay (IFA), immunoenzymatic assay (ELISA), and the immunochromatographic test (IC) are also frequently utilized for the diagnosis of leishmaniasis. Other less employed serological tests include direct agglutination test (DAT), fast agglutination screening test (FAST), the complement fixation reaction (CFR), and western blot (WB). The sensitivity and specificity of these methods are directly related to the technique used and the manifestation of the disease [ 11 , 19 ]. Moreover, these antibody detection tests do not differentiate between recent and past infections, as the antibodies remain positive months after curing the patient [ 14 ]. The IFA test shows acceptable sensitivity (87–100%) and specificity (77–100%) [ 8 , 20 , 21 ]. However, the need for a fluorescence microscope limits the use of the IFA test to reference laboratories [ 14 ]. ELISA is the preferred laboratory test for serodiagnosis of VL. The technique is highly sensitive, but its specificity depends upon the antigens used. Variations in sensitivity (80–100%) and specificity (71–100%) are reported in ELISA tests using crude Leishmania spp. antigens [ 22 – 24 ]. The use of recombinant or purified antigens, such as membrane glycoproteins gp63, gp72, gp70, and rK39 specific to the Leishmania genus, improve the sensitivity and specificity of the technique. However, cross-reactions with diseases caused by other trypanosomatids can still occur [ 23 ]. In view of the need to achieve high precision, several new serological methodologies have been developed in recent years, and research using rapid immunochromatographic tests has been explored. Among these, the point of care (POC) tests using different antigens are popular, however they show varying sensitivity and specificity among different populations [ 14 ]. With respect to cytometry, although the technique has also shown high sensitivity and specificity, and presents advantages over other immunoassays, it is costly and requires specialized technicians to manipulate the equipment. Thus, its use in diagnostic routine is limited and is usually intended for large research centers [25–27]. Pharmaceutics 2021,13, 491 4 of 19 2.3. Molecular Tests Polymerase chain reaction (PCR) is a highly sensitive and important method used for the diagnosis of leishmaniasis, mainly in immunosuppressed patients. PCR can be performed using a wide variety of samples, such as blood, spleen, and lymph node [ 10 , 14 ]. One of its main advantages is the possibility of quantifying the parasitic load in the samples (qPCR), allowing the progression of the disease to be monitored and, consequently, the effectiveness of the anti-Leishmania therapy used [ 10 , 14 ]. Molecular techniques, although not used in the routine diagnosis, are highly sensitive. PCR has a reported specificity of 100% (for CL) with an improved sensitivity of 20 to 30% when compared with conventional parasitology diagnosis, and thus its inclusion as one of the routinely diagnostic methods could be highly beneficial. Real-time quantitative PCR (qPCR) has been shown to have high analytical sensitivity (0.0125 parasites per mL of blood) and excellent linearity [ 28 ]. PCR allows identification of the species of Leishmania that is causing the infection [ 10 , 29 ]. However, the lack of standardization in the protocols for the different reaction steps (obtaining the samples, extracting the DNA, and selecting the primers) makes it difficult to implement the technique on a large scale [ 11 , 30 ]. Furthermore, these tests remain limited to reference hospitals and research centers because of high costs and the requirement for special equipment and skilled personnel. Efforts are needed to make PCR more economical and easier to use, especially in endemic areas [ 11 , 14 ]. Despite the great progress, there is still no standard test for the diagnosis of leishmaniasis. An effective diagnostic method, needs to be sensitive, specific, fast, accessible, and easy to use, among other factors. Together, these factors will result in early efficient disease identification. 3. Emerging Nanomaterial-Based Detection Technologies The limitations encountered in the methods currently in use for the diagnosis of Leishmania demand development of new, efficient, rapid, and innovative POC approaches. Over the years, biosensors and nano-based technologies have proven to be promising options to meet this ever-increasing demand. Biosensors consist of a biomolecule (antigen, antibody, oligonucleotide, or enzyme) as recognition element coupled to a signal transducer. Binding of the recognition element to the target generates a signal by the transducer [ 31 ]. Based on the recognition element, biosensors can be categorized as immunosensors, genosensors, and aptamer-based sensors [ 32 ]. Immunosensors use antibodies as the recognition element. The transducer in this case converts the antibody–antigen interaction into a measurable physical signal. Genosensors, on the other hand, are DNA biosensors involving a hybridization reaction between two complementary oligonucleotides [ 33 ]. In this case, the use of nucleic acids eliminates the need for expensive antibodies, resulting in higher stability and discarding the need for special storage. Aptamer-based sensors make use of aptamers (single-stranded DNA oligonucleotides that can specifically bind target analytes) also known as chemical antibodies [34]. Nanomaterials are known for their excellent optical, catalytic, electrical, and magnetic properties [ 35 – 39 ]. Many published reports describe the use of these special physical and chemical characteristics of nanomaterials for the development of biosensors and other nano-based tools for the detection of Leishmania which will be described in this section (Table 1). Table 1. Nano-based biosensors described for the detection of Leishmania. Type of Biosensor Nanomaterial Used Type of Pathogen Target Biomolecule Limit of Detection References Genosensor/Optical PPY PANI L. infantum FAM-ssDNA 1.1 nM † 1.3 nM [23] Genosensor/Optical cadmium selenite QD Leishmania spp. Conserved specific genomic DNA LPG and gp36 antigens 3.125 ng/µL‡ [40] Immunosensor/Optical 103cells/mL ¥ Pharmaceutics 2021,13, 491 5 of 19 Table 1. Cont. Type of Biosensor Nanomaterial Used Type of Pathogen Target Biomolecule Limit of Detection References Genosensor/Optical AuNP L. infantum KinetoplastDNA 100 fmol [41] Genosensor/Optical AuNP L. major KinetoplastDNA 7.0 pg/µL [42] Genosensor/Optical AuNP Leishmania spp. KinetoplastDNA 11.5 ng/µL [43] Immunosensor/Optical AuNP L. infantum Chimeric recombinant antigens (K9, K39, and K26) - [44] Immunosensor/Impedimetric PAMAM L. amazonensis membrane proteins 10−5mg/mL [45] Immunosensor/SPR PAMAM L. infantum hypothetical C1protein 7.83 nmol/L [46] Genosensor/Electrochemical cobalt-zinc ferrite QD L. major KinetoplastDNA 1.8 ×10−14 ng/µL[47] Genosensor/Electrochemical AuNP L. major L. major specific DNA 1.8 ×10−20 mol/L 0.7 ng/µL[48] Genosensor/Electrochemical AuNP L. infantum KinetoplastDNA 2×10−19 mol/L [49] Genosensor/Electrochemical AuNP Leishmania spp. Leishmania specific DNA 1ZM [50] Casein-gp63 interaction/Electrochemical AuNP L. infantum gp63 surface protein 0.55 parasite/mL [51] Genosensor/Electrochemical AuNP Leishmania spp. KinetoplastDNA 0.8 parasite/mL [52] Aptasensor/Electrochemical AuNP L. infantum Kinetoplastid membrane protein-11 2.27 mM [53] PPY—polypyrrole, PAN—polyaniline, QD—quantum dot, PAMAM—polyamidoamine dendrimer, AuNP—gold nanoparticle, FAMssDNA—6-carboxyfluorescein-labeled single stranded DNA; † , In this case, the value represents the concentration of DNA as the target molecule is ssDNA from L. infantum; ‡ In this case, the value represents the concentration of DNA as target molecule is genomic DNA; ¥ , In this case, the value represents the number of parasites. 3.1. Nanomaterials as Fluorescence Quenchers Various types of nanostructures such carbon nanotubes; graphene oxide [ 54 , 55 ]; Au [ 56 ]; Ag [ 57 , 58 ] and ceria [ 59 ] nanoparticles; quantum dots [ 60 ]; metal–organic structures [ 61 , 62 ]; transition metal nanosheets [ 63 , 64 ]; and conductive polymers—such as poly(3,4-ethylenedioxythiophene) (PEDOT) nanoparticles [ 65 ], polypyrrole (PPY) nanospheres [ 66 ], and polyaniline (PANI) nanofibers, have been reported as effective fluorescence quenchers. Immobilizing these nanostructures on flexible substrates and their further integration in portable devices have been explored in many different applications. Functionalized nanostructures (with the probe of interest) have been used as fluorescence test strips, reducing the time required to carry out a diagnostic assay. Pedro et al. (2019) [ 67 ] described the use of nanomaterials as nanoquenchers for fluorescent DNA assays. The nanostructure developed interacted with the fluorophore of a labeled DNA probe (L. infantum specific) through electron transfer processes which resulted in quenching of the fluorescence emission. Restoration of the fluorescence could only be achieved in the presence of complementary DNA to the original DNA probe. Subsequently, the probe–target hybridization resulted in desorption of the labeled probe into the supernatant. In their work, the authors used nanostructured ICP/PET (intrinsically conducting polymers/polyethylene terephthalate) films for the detection of L. infantum specific DNA sequences (Figure 2). The DNA detection system utilized PPY and PANI films immobilized with 6-carboxyfluorescein-labeled single stranded DNA (FAM-ssDNA), resulting in the quenching of the luminescence signal. Later, when the FAM-ssDNA/ICP film system was put in contact with the target ssDNA, the hybridization of the probe with its complementary target caused the formation of double-stranded DNA, release of the FAM-ssDNA from the surface of the films, and fluorescence recovery. Both types of polymeric films utilized in the study showed high sensitivity, with respective detection limits of 1.1 nM (PPY) and 1.3 nM (PANI) against the target ssDNA. Furthermore, the Pharmaceutics 2021,13, 491 6 of 19 authors reported a linear response for the fluorescence intensity in a large concentration range of the target DNA (1–300 nM and 1–500 nM, for PPY and PANI, respectively). The FAM-ssDNA/ICP sensors showed high selectivity and could differentiate between full complementary sequences and those with just a single mismatch. They could also function in spiked complex medium, such as human blood serum, and exhibited good shelf stability. The functionalized ICP films also exhibited good shelf stability (PPY/PANI films were functional up to 180 days and probe immobilized films were functional for several days). The nanostructured ICP films described in this study show potential practical applications as strip tests for fast and sensitive fluorescent DNA detection in point-of-care clinical diagnosis protocols. This study thus describes the development of an inexpensive and simple molecular test that would not necessarily replace PCR analyses, but would rather complement them as efficient pre-screening methods. In this method, although PCR amplification is not required, both DNA extraction from a biological sample and the conversion of double-stranded DNA (dsDNA) to ssDNA are still necessary. Pharmaceutics 2021, 13, x FOR PEER REVIEW 6 of 20 films were functional up to 180 days and probe immobilized films were functional for several days). The nanostructured ICP films described in this study show potential practical applications as strip tests for fast and sensitive fluorescent DNA detection in pointof-care clinical diagnosis protocols. This study thus describes the development of an inexpensive and simple molecular test that would not necessarily replace PCR analyses, but would rather complement them as efficient pre-screening methods. In this method, although PCR amplification is not required, both DNA extraction from a biological sample and the conversion of double-stranded DNA (dsDNA) to ssDNA are still necessary. . Figure 2. Nanostructured films as quenchers for fluorescent detection. Mechanism of fluorescence detection (of nucleic acids) using nanostructured ICP/PET films as sensing platform is demonstrated. 3.2. Nanomaterial as Fluorophore Quantum dots (QDs) are nanosized semiconducting materials that have proven to be 20 times brighter and 100 times more stable compared to conventional fluorescent probes. Andreadou et al. (2016) [40] showed the use of cadmium selenite (CdSe) QDs as signal detector for the detection of Leishmania. Two different systems were described by this group: one for detection of Leishmania-specific DNA, and another for the detection of Leishmania-specific surface proteins (LPG and gp63). In both, the system’s magnetic beads were used to separate the analytes from the solution, whereas the presence of the targeted molecules was demonstrated by using QDs. The aim here was to develop a methodology that would be technically easy to perform, not require a specific equipment, and at the same time detect all the main pathogens of Leishmania spp. For the DNA-based biosensor, two highly specific DNA oligonucleotides (probe 1 and probe 2) conserved among the main leishmanial pathogens were utilized as DNA probes [68]. These DNA probes (probes 1 and 2) were biotinylated at their 5’ end and detected with streptavidin-bound QDs (15–20 nm in size). Denatured single-stranded target DNA was first put in contact with biotinylated probe 1 and was separated using the magnetic beads conjugated with streptavidin. The separated mixture was incubated with biotinylated probe 2 and finally detected with QDs bound to streptavidin (Figure 3A). A similar methodology was used for the detection of the two different leishmanial surface antigens (LPG and gp63), using monoclonal antibodies, biotinylated IgG/IgM, and streptavidin-bound QDs. Magnetic beads conjugated with streptavidin were used as separation probe (Figure 3B). The fluorescent Figure 2. Nanostructured films as quenchers for fluorescent detection. Mechanism of fluorescence detection (of nucleic acids) using nanostructured ICP/PET films as sensing platform is demonstrated. 3.2. Nanomaterial as Fluorophore Quantum dots (QDs) are nanosized semiconducting materials that have proven to be 20 times brighter and 100 times more stable compared to conventional fluorescent probes. Andreadou et al. (2016) [ 40 ] showed the use of cadmium selenite (CdSe) QDs as signal detector for the detection of Leishmania. Two different systems were described by this group: one for detection of Leishmania-specific DNA, and another for the detection of Leishmania-specific surface proteins (LPG and gp63). In both, the system’s magnetic beads were used to separate the analytes from the solution, whereas the presence of the targeted molecules was demonstrated by using QDs. The aim here was to develop a methodology that would be technically easy to perform, not require a specific equipment, and at the same time detect all the main pathogens of Leishmania spp. For the DNA-based biosensor, two highly specific DNA oligonucleotides (probe 1 and probe 2) conserved among the main leishmanial pathogens were utilized as DNA probes [ 68 ]. These DNA probes (probes 1 and 2) were biotinylated at their 5’ end and detected with streptavidin-bound QDs (15–20 nm Pharmaceutics 2021,13, 491 7 of 19 in size). Denatured single-stranded target DNA was first put in contact with biotinylated probe 1 and was separated using the magnetic beads conjugated with streptavidin. The separated mixture was incubated with biotinylated probe 2 and finally detected with QDs bound to streptavidin (Figure 3A). A similar methodology was used for the detection of the two different leishmanial surface antigens (LPG and gp63), using monoclonal antibodies, biotinylated IgG/IgM, and streptavidin-bound QDs. Magnetic beads conjugated with streptavidin were used as separation probe (Figure 3B). The fluorescent signal generated in both the systems was detected visually or by optical analysis (605 nm). Positive results were obtained in all the positive samples analyzed, and no fluorescent signal was obtained in the negative control samples, thus generating a sensitivity and specificity of 100%. The limit of detection was calculated to be 3.125 ng/ µ L and 10 3 cells/mL for the DNA and protein methods, respectively. Pharmaceutics 2021, 13, x FOR PEER REVIEW 7 of 20 signal generated in both the systems was detected visually or by optical analysis (605 nm). Positive results were obtained in all the positive samples analyzed, and no fluorescent signal was obtained in the negative control samples, thus generating a sensitivity and specificity of 100%. The limit of detection was calculated to be 3.125 ng/μL and 10 3 cells/mL for the DNA and protein methods, respectively. Figure 3. (A) Detection of Leishmania-specific target DNA using quantum dots. Target DNA was first denatured, followed by the hybridization of the functionalized magnetic beads to quantum dots via complimentary sequences. A magnetic device was then used to separate the conjugates from the solution and the positive result was detected by fluorescence. (B) Use of quantum dots for detecting Leishmania-specific proteins. Leishmanial cells were first conjugated with a monoclonal anti-Leishmania LPG antibody which initially bound to a biotin-labeled polyclonal anti-mouse IgM and later to streptavidin-coated magnetic beads. After separation using a magnet, the second monoclonal antibody (anti-Leishmania gp63) first bound to the protozoan cells and then to the biotin labeled polyclonal anti-mouse IgG. The complexes were separated again using a magnet and finally detected by streptavidin-coated quantum dots. 3.3. Use of Dendrimers Perinoto et al. (2010) [45] developed a nanostructured biosensor system to detect specific anti-Leishmania antibodies using capacitance measurements. In this system, phospholipid liposomes incorporating L. amazonensis-specific membrane antigenic proteins (proteolisosomes) together with dendrimers attached to the surface of interdigitated gold electrodes were used as the immobilized phase. The electrodes containing antigenic proteins were then used to detect antibodies. Binding of the antibodies to the electrode resulted in variations in the measured electrical response. Freshly prepared proteoliposomes were immobilized on interdigitated electrodes using the layer-by-layer technique in conjunction with polyamidoamine dendrimers 4 (PAMAM). Dendrimers, due to their branched, porous structure, are utilized for immobilizing proteins in biosensors, as they increase Figure 3. ( A ) Detection of Leishmania-specific target DNA using quantum dots. Target DNA was first denatured, followed by the hybridization of the functionalized magnetic beads to quantum dots via complimentary sequences. A magnetic device was then used to separate the conjugates from the solution and the positive result was detected by fluorescence. ( B ) Use of quantum dots for detecting Leishmania-specific proteins. Leishmanial cells were first conjugated with a monoclonal antiLeishmania LPG antibody which initially bound to a biotin-labeled polyclonal anti-mouse IgM and later to streptavidin-coated magnetic beads. After separation using a magnet, the second monoclonal antibody (anti-Leishmania gp63) first bound to the protozoan cells and then to the biotin labeled polyclonal anti-mouse IgG. The complexes were separated again using a magnet and finally detected by streptavidin-coated quantum dots. 3.3. Use of Dendrimers Perinoto et al. (2010) [ 45 ] developed a nanostructured biosensor system to detect specific anti-Leishmania antibodies using capacitance measurements. In this system, phospholipid liposomes incorporating L. amazonensis-specific membrane antigenic proteins (proteolisosomes) together with dendrimers attached to the surface of interdigitated gold Pharmaceutics 2021,13, 491 8 of 19 electrodes were used as the immobilized phase. The electrodes containing antigenic proteins were then used to detect antibodies. Binding of the antibodies to the electrode resulted in variations in the measured electrical response. Freshly prepared proteoliposomes were immobilized on interdigitated electrodes using the layer-by-layer technique in conjunction with polyamidoamine dendrimers 4 (PAMAM). Dendrimers, due to their branched, porous structure, are utilized for immobilizing proteins in biosensors, as they increase sensitivity and response times due to diffusion of analytes through the multilayer structure. Furthermore, the porous architecture of the film is important for confining electrical charges within its structure, which are responsible for the detectable changes in the electrical response. The PAMAM/proteoliposome electrode developed in this study could differentiate the capacitance signal between the positive and negative serum samples without showing any cross-reactivity with anti-T. cruzi IgGs, which are responsible for significant false positive in the current diagnostic methodologies. The capacitance-based system developed here can be extended for the diagnosis of other bacterial, protozoan, and helminth infectious diseases. Souto et al. (2015) [ 46 ] also made use of PAMAM for detection of anti-L. infantum antibodies using a surface plasmon resonance (SPR)-based immunosensor. The authors used recombinant protein from L. infantum with unknown function (hypothetical C1 protein or C1 antigen) as a recognition element for the development of SPR immunosensor. SPRbased sensors are widely recognized as a potential analytical tool due to their extreme sensitivity to small changes in the refractive index near to the sensor surface caused by the variation of the mass on the transducer surface. SPR requires neither label nor tracer, thus reducing the number of steps for the analysis of biomolecular interactions in real time. In the current study, the immunosensor was developed by depositing selfassembled layers (SAM) of cysteamine on gold surface followed by the addition of a fourthgeneration poly (amidoamine) dendrimer (PAMAM (G4)) onto which the C1 antigen was immobilized. Binding of antibody to the immobilized antigen resulted in the variation of the angle of resonance which was recorded. The authors in this work confirmed the specific binding of CVL (canine visceral leishmania) antibodies to the immobilized recombinant C1 antigen showing a limit of detection of 7.83 nmol/L. No signal was obtained for negative canine sera. A similar result was also shown by this group using different L. infantum antigens immobilized on SAM of 11-MUA on gold substrate for the development of a SPR immunosensor in canine serum [69]. 3.4. Surface Plasmon Coupling Detection of the amplification products (e.g., gel electrophoresis) after PCR amplification requires time-consuming protocols performed by trained personnel, with high cost. The aim of the study conducted by Toubanaki et al. (2016) [ 41 ] was simplification of PCR product detection, using a nucleic acid lateral flow, combined with functionalized gold nanoparticles. Gold nanoparticles, besides providing high specific surface (thus enabling the immobilization of an increased amount of bioreceptor units), were also used for visualization based on their interparticle surface plasmon coupling [ 70 , 71 ]. Lateral flow biosensors (LFBs) are diagnostic devices based on one time use of paper as a carrier material, where dry reagents are activated by applying a fluid sample. LFBs are important for diagnostic purposes as they are affordable, sensitive, specific, user-friendly, rapid, robust, and equipment-free [ 71 , 72 ]. They are an ideal platform for single use at point of care, where only a positive/negative signal is desired, with fast result time (approximately 20 min) [ 73 , 74 ]. In the current study [ 41 ], amplification reactions targeting kinetoplast DNA of Leishmania spp. Were performed on canine blood samples, and a positive signal was obtained as a red test zone on LFB. LFB consisted of an immersion pad (17 mm), a glassfiber conjugate pad (8 mm), nitrocellulose diagnostic membrane (25 mm in length), and an absorbent pad (same as immersion pad) assembled on a plastic adhesive backing. The genomic DNA extracted from the canine sample was subjected to PCR for kinetoplast DNA amplification using biotinylated primers. Amplified product was mixed with dATP-tailed Leishmania-specific probe, and the mixture was applied to the conjugation pad next to the Pharmaceutics 2021,13, 491 9 of 19 poly (dT) conjugated gold. Gold nanoparticles functionalized with oligo (dT) segments interacted with the hybridized complex of target DNA–poly (dA) probe and then were captured by immobilized streptavidin in the test zone of the LFB, indicating a positive signal (characteristic red line due to gold nanoparticle accumulation). The visual detection was completed in 20 min. Extensive optimization enabled the detection of 100 mol of target DNA. The biosensor was evaluated with actual clinical samples of infected dog blood and successfully confirmed the presence of Leishmania, while no product was detected for negative samples. Overall, the proposed lateral flow biosensor described by the authors is an appealing alternative platform for the detection of Leishmania-specific amplification products with low cost and attractive simplicity. Detection of the PCR products via LFB provides sequence confirmation by target–probe hybridization compared to size-based recognition of the amplified fragments in case of electrophoresis. The pre-hybridized sample can be applied directly on the biosensor, and only a small amount of developing solution and gold nanoparticles is required for visualization, eliminating the need of gel electrophoresis. The cost of the assay in terms of reagents and the biosensor was described to be about three euro. Furthermore, the biosensor allowed visual detection of Leishmania-specific amplification products within minutes without the need of any instruments. Overall, the proposed lateral flow biosensor can be considered an appealing alternative platform for Leishmania-specific amplification products detection with low cost and attractive simplicity. Sattarahmady et al. (2016) [ 42 ] reported a gold nanoparticle-based genosensor for the visual and spectrophotometric detection of non-protein-coding region of the kDNA minicircle genome from L. major. The mechanism was based on hybridization of AuNPsprobeDNA with a target complementary DNA sequence. Changes in the plasmonic property of AuNPs, which depends on size, surface function, and interval distance between the particles, were used to induce a color change which was used for sensing. AuNPs (16 nm) used in this study had a stable red color which turned to purple on the addition of HCl due to aggregation by acid. However, no color change was observed in the presence of complimentary sequences, probably due to the formation of double-stranded DNA, which increases the stability of the AuNP probe toward aggregation. The method described by the authors showed specificity for detection of L. major with a detection limit of 7.0 pg/ µ L. The method was evaluated with success for the detection of L. major genomic DNA (DNA extracts of standard cultures of L. major) as well as its detection in clinical samples (DNA extracted from clinical samples). Based on the same principles (AuNP and aggregation by HCl), Andreadou et al. (2014) [ 43 ] designed a genosensor using four single-stranded oligonucleotides based on conserved regions of Leishmania genome as probe and confirmed the specificity and sensitivity of the sensor using positive and negative control samples and the whole blood collected from dogs with suspected canine leishmaniasis. The minimum detection limit was defined to 11.5 ng/ µ L of target DNA sample with 100% repeatability and reproducibility. Anfossi et al. (2018) [ 44 ] also described a lateral flow immunoassay (LFIA) for quick diagnosis of canine leishmaniasis which can also be extended to other animals (Figure 4). The device used highly specific chimeric recombinant antigens (K9, K39, and K26) from the amastigote form of L. infantum and measured anti-leishmanial antibodies present in the serum. Protein A (from Staphylococcus known to bind to immunoglobins from various animals) labeled with gold nanoparticles (30 nm mean diameter and a SPR band at 525 nm) was used as the signal reporter, and the visual results after 15 minutes were comparable to that generated by ELISA and IFAT. The developed LFIA showed high diagnostic sensitivity (98.4%) and specificity (98.9%), in agreement with serological reference methods for diagnosing canine visceral leishmaniasis. Furthermore, the authors confirmed long thermal stability (six months of storage at room temperature or 4 ◦C). Pharmaceutics 2021,13, 491 16 of 19 amplification has also been well explored. However, further improvements are required with respect to the specificity of the transduced signal evicting cross-reactions. The use of conducting, catalytic, optical, and structural properties of nanomaterials are promising; however, the question is: Is this technique is viable to be used at large scale with an effective cost–benefit ratio? Most of the recognition elements in biosensors described in this review showed the use of antibodies with only one study making use of aptamers. Aptamers are highly promising with superior properties compared to antibodies. Selection of new, more sensitive aptamers with better and more systematic characterization is definitely desired. Moreover, most of the aptamer-based biosensors rely on simple binding and are prone to nonspecific binding, especially in a complex sample matrix. An electrochemical DNA (E-DNA) sensor platform could be a good alternative, where signaling does not rely on simple absorption of the target on the sensor, thus the system performs well even in a complex medium. Sensors based on aptamer conformational change or catalysis could thus be more accurate. Author Contributions: All authors have substantially contributed to the conceptualization, methodology, formal analysis of collected data, building up of figures and tables, investigation, resources, and data analysis. S.J., W.S., S.S.D., and A.L.S.S. have contributed to the writing—original draft, data search, and literature research. S.J., E.B.S., and P.S. have contributed for the writing—review, validation, and editing. S.J., W.S., S.S.D., A.L.S.S., E.B.S., and P.S. have contributed to the project administration, supervision, and funding acquisition. All authors have read and agreed to the published version of the manuscript. Funding: This research was funded by Conselho Nacional de Desenvolvimento Científico e Tecnológico for the scientific grants (CNPq 301964/2019-0 Chamada 06/2019, Chamada CNPq n º 01/2019) and Portuguese Science and Technology Foundation, Ministry of Science and Education (FCT/MEC) through the sponsorship of the projects M-ERA-NET-0004/2015-PAIRED and UIDB/04469/2020 (strategic fund), co-financed by FEDER, under the Partnership Agreement PT2020. Institutional Review Board Statement: Not applicable. Informed Consent Statement: Not applicable. Data Availability Statement: Not applicable. Conflicts of Interest: The authors declare no conflict of interest. References 1. Torres-Guerrero, E.; Quintanilla-Cedillo, M.R.; Ruiz-Esmenjaud, J.; Arenas, R. Leishmaniasis: A review. F1000Research 2017 , 6, 750. [CrossRef] 2. De las Heras, F.G. Overview of Neglected Tropical Diseases. In Third World Diseases, Elliott, R., Ed.; Springer: Berlin/Heidelberg, Germany, 2011; pp. 1–46. [CrossRef] 3. Burza, S.; Croft, S.L.; Boelaert, M. Leishmaniasis. Lancet 2018,392, 951–970. [CrossRef] 4. Herrera, G.; Barragán, N.; Luna, N.; Martínez, D.; De Martino, F.; Medina, J.; Niño, S.; Páez, L.; Ramírez, A.; Vega, L.; et al. An interactive database of Leishmania species distribution in the Americas. Sci. Data 2020,7, 110. [CrossRef] [PubMed] 5. Alvar, J.; Vélez, I.D.; Bern, C.; Herrero, M.; Desjeux, P.; Cano, J.; Jannin, J.; Den Boer, M. Leishmaniasis worldwide and global estimates of its incidence. PLoS ONE 2012,7, e035671. [CrossRef] [PubMed] 6. Quinnell, R.J.; Courtenay, O. Transmission, reservoir hosts and control of zoonotic visceral leishmaniasis. Parasitology 2009 , 136, 1915–1934. [CrossRef] [PubMed] 7. Aronson, N.; Herwaldt, B.L.; Libman, M.; Pearson, R.; Lopez-Velez, R.; Weina, P.; Carvalho, E.; Ephros, M.; Jeronimo, S.; Magill, A. Diagnosis and Treatment of Leishmaniasis: Clinical Practice Guidelines by the Infectious Diseases Society of America (IDSA) and the American Society of Tropical Medicine and Hygiene (ASTMH). Am. J. Trop. Med. Hyg. 2017,96, 24–45. [CrossRef] 8. McGwire, B.S.; Satoskar, A.R. Leishmaniasis: Clinical syndromes and treatment. QJM Int. J. Med. 2013,107, 7–14. [CrossRef] 9. Varma, N.; Naseem, S. Hematologic changes in visceral leishmaniasis/kala azar. Indian J. Hematol. Blood Transfus. 2010 , 26, 78–82. [CrossRef] 10. Akhoundi, M.; Downing, T.; Votýpka, J.; Kuhls, K.; Lukeš, J.; Cannet, A.; Ravel, C.; Marty, P.; Delaunay, P.; Kasbari, M.; et al. Leishmania infections: Molecular targets and diagnosis. Mol. Asp. Med. 2017,57, 1–29. [CrossRef] 11. Roatt, B.M.; De Oliveira Cardoso, J.M.; De Brito, R.C.F.; Coura-Vital, W.; De Oliveira Aguiar-Soares, R.D.; Reis, A.B. Recent advances and new strategies on leishmaniasis treatment. Appl. Microbiol. Biotechnol. 2020,104, 8965–8977. [CrossRef] Pharmaceutics 2021,13, 491 17 of 19 12. Goto, H.; Lindoso, J.A. Current diagnosis and treatment of cutaneous and mucocutaneous leishmaniasis. Expert Rev. Anti-Infect. Ther. 2010,8, 419–433. [CrossRef] [PubMed] 13. Georgiadou, S.P.; Makaritsis, K.P.; Dalekos, G.N. Leishmaniasis revisited: Current aspects on epidemiology, diagnosis and treatment. J. Transl. Int. Med. 2015,3, 43–50. [CrossRef] 14. Kotb, E.E.; Antonio Sampedro, M.; Javier, R.-G.; Yannick, H.-M.; Ahamd, A.; Jose Mari Navarro, M.; Jose Gutierrez, F. Diagnosis of leishmaniasis. J. Infect. Dev. Ctries. 2014,8. [CrossRef] 15. Liu, J.; Cao, Z.; Lu, Y. Functional Nucleic Acid Sensors. Chem. Rev. 2009,109, 1948–1998. [CrossRef] [PubMed] 16. Alizadeh, N.; Memar, M.Y.; Moaddab, S.R.; Kafil, H.S. Aptamer-assisted novel technologies for detecting bacterial pathogens. Biomed. Pharmacother. 2017,93, 737–745. [CrossRef] [PubMed] 17. Majdinasab, M.; Hayat, A.; Marty, J.L. Aptamer-based assays and aptasensors for detection of pathogenic bacteria in food samples. TrAC Trends Anal. Chem. 2018,107, 60–77. [CrossRef] 18. Blum, J.; Desjeux, P.; Schwartz, E.; Beck, B.; Hatz, C. Treatment of cutaneous leishmaniasis among travellers. J. Antimicrob. Chemother. 2004,53, 158–166. [CrossRef] 19. Molina, I.; Fisa, R.; Riera, C.; Falcó, V.; Elizalde, A.; Salvador, F.; Crespo, M.; Curran, A.; López-Chejade, P.; Tebar, S.; et al. Ultrasensitive real-time PCR for the clinical management of visceral leishmaniasis in HIV-Infected patients. Am. J. Trop. Med. Hyg. 2013,89, 105–110. [CrossRef] 20. Iqbal, J.; Hira, P.R.; Saroj, G.; Philip, R.; Al-Ali, F.; Madda, P.J.; Sher, A. Imported visceral leishmaniasis: Diagnostic dilemmas and comparative analysis of three assays. J. Clin. Microbiol. 2002,40, 475–479. [CrossRef] [PubMed] 21. Boelaert, M.; Rijal, S.; Regmi, S.; Singh, R.; Karki, B.; Jacquet, D.; Chappuis, F.; Campino, L.; Desjeux, P.; Le Ray, D.; et al. A comparative study of the effectiveness of diagnostic tests for visceral leishmaniasis. Am. J. Trop. Med. Hyg. 2004 ,70, 72–77. [CrossRef] [PubMed] 22. Sundar, S.; Rai, M. Laboratory Diagnosis of Visceral Leishmaniasis. Clin. Diagn. Lab. Immunol. 2002,9, 951–958. [CrossRef] [PubMed] 23. Pedras, M.J.; De Gouvêa Viana, L.; De Oliveira, E.J.; Rabello, A. Comparative evaluation of direct agglutination test, rK39 and soluble antigen ELISA and IFAT for the diagnosis of visceral leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 2008 ,102, 172–178. [CrossRef] [PubMed] 24. De Assis, T.S.M.; Caligiorne, R.B.; Romero, G.A.S.; Rabello, A. Detection of Leishmania kDNA in human serum samples for the diagnosis of visceral leishmaniasis. Trans. R. Soc. Trop. Med. Hyg. 2009,103, 1269–1272. [CrossRef] [PubMed] 25. Fagundes, A.; Schubach, A.; Paula, C.C.d.; Bogio, A.; Antonio, L.d.F.; Schiavoni, P.B.; Monteiro, V.d.S.; Madeira, M.d.F.; Quintella, L.P.; Valete-Rosalino, C.M.; et al. Evaluation of polymerase chain reaction in the routine diagnosis for tegumentary leishmaniasis in a referral centre. J. Memórias Inst. Oswaldo Cruz. 2010,105, 109–112. [CrossRef] [PubMed] 26. De Castro Zacche-Tonini, A.; Fonseca, G.S.F.; De Jesus, L.; Barros, G.B.; Coelho-Dos-Reis, J.G.A.; Béla, S.R.; Machado, A.S.; Carneiro, A.; Andrade, G.M.Q.; Vasconcelos-Santos, D.V.; et al. Establishing tools for early diagnosis of congenital toxoplasmosis: Flow cytometric IgG avidity assay as a confirmatory test for neonatal screening. J. Immunol. Methods 2017 ,451, 37–47. [CrossRef] 27. Alessio, G.D.; De Araújo, F.F.; Sales Júnior, P.A.; Gomes, M.S.; Amaral, L.R.D.; Pascoal Xavier, M.A.; Teixeira-Carvalho, A.; De Lana, M.; Martins-Filho, O.A. Accomplishing the genotype-specific serodiagnosis of single and dual Trypanosoma cruzi infections by flow cytometry Chagas-Flow ATE-IgG2a. PLoS Negl. Trop. Dis. 2018,12, e0006140. [CrossRef] 28. Mary, C.; Faraut, F.; Lascombe, L.; Dumon, H. Quantification of Leishmania infantum DNA by a real-time PCR assay with high sensitivity. J. Clin. Microbiol. 2004,42, 5249–5255. [CrossRef] 29. Schönian, G.; Kuhls, K.; Mauricio, I.L. Molecular approaches for a better understanding of the epidemiology and population genetics of Leishmania. Parasitology 2011,138, 405–425. [CrossRef] 30. Hossain, F.; Ghosh, P.; Khan, M.A.A.; Duthie, M.S.; Vallur, A.C.; Picone, A.; Howard, R.F.; Reed, S.G.; Mondal, D. Real-time PCR in detection and quantitation of Leishmania donovani for the diagnosis of Visceral Leishmaniasis patients and the monitoring of their response to treatment. PLoS ONE 2017,12, e0185606. [CrossRef] 31. Bhalla, N.; Jolly, P.; Formisano, N.; Estrela, P. Introduction to biosensors. Essays Biochem. 2016,60, 1–8. [CrossRef] 32. Zhang, Z.; Cong, Y.; Huang, Y.; Du, X. Nanomaterials-Based Electrochemical Immunosensors. Micromachines 2019 ,10, 397. [CrossRef] [PubMed] 33. Kaushik, M.; Khurana, S.; Mehra, K.; Yadav, N.; Mishra, S.; Kukreti, S. Emerging Trends in Advanced Nanomaterials Based Electrochemical Genosensors. Curr. Pharm. Des. 2018,24, 3697–3709. [CrossRef] [PubMed] 34. Yu, X.; Yang, Y.-P.; Dikici, E.; Deo, S.K.; Daunert, S. Beyond Antibodies as Binding Partners: The Role of Antibody Mimetics in Bioanalysis. Annu. Rev. Anal. Chem. 2017,10, 293–320. [CrossRef] 35. Kumar, S.; Ahlawat, W.; Kumar, R.; Dilbaghi, N. Graphene, carbon nanotubes, zinc oxide and gold as elite nanomaterials for fabrication of biosensors for healthcare. Biosens. Bioelectron. 2015,70, 498–503. [CrossRef] 36. Shavanova, K.; Bakakina, Y.; Burkova, I.; Shtepliuk, I.; Viter, R.; Ubelis, A.; Beni, V.; Starodub, N.; Yakimova, R.; Khranovskyy, V. Application of 2D Non-Graphene Materials and 2D Oxide Nanostructures for Biosensing Technology. Sensors 2016 ,16, 223. [CrossRef] [PubMed] 37. Hu, Y.; Cheng, H.; Zhao, X.; Wu, J.; Muhammad, F.; Lin, S.; He, J.; Zhou, L.; Zhang, C.; Deng, Y.; et al. Surface-Enhanced Raman Scattering Active Gold Nanoparticles with Enzyme-Mimicking Activities for Measuring Glucose and Lactate in Living Tissues. ACS Nano 2017,11, 5558–5566. [CrossRef] Pharmaceutics 2021,13, 491 18 of 19 38. Jain, S.; Huang, Z.; Dixon, B.R.; Sattar, S.; Liu, J. Cryptosporidium parvum oocyst directed assembly of gold nanoparticles and graphene oxide. Front. Chem. Sci. Eng. 2019,13, 608–615. [CrossRef] 39. Jain, S.; Costa Melo, T.G.; Dolabella, S.S.; Liu, J. Current and emerging tools for detecting protozoan cysts and oocysts in water. TrAC Trends Anal. Chem. 2019,121, 115695. [CrossRef] 40. Andreadou, M.; Liandris, E.; Gazouli, M.; Mataragka, A.; Tachtsidis, I.; Goutas, N.; Vlachodimitropoulos, D.; Ikonomopoulos, J. Detection of Leishmania-specific DNA and surface antigens using a combination of functionalized magnetic beads and cadmium selenite quantum dots. J. Microbiol. Methods 2016,123, 62–67. [CrossRef] 41. Toubanaki, D.K.; Athanasiou, E.; Karagouni, E. Gold nanoparticle-based lateral flow biosensor for rapid visual detection of Leishmania-specific DNA amplification products. J. Microbiol. Methods 2016,127, 51–58. [CrossRef] [PubMed] 42. Sattarahmady, N.; Movahedpour, A.; Heli, H.; Hatam, G.R. Gold nanoparticles-based biosensing of Leishmania major kDNA genome: Visual and spectrophotometric detections. Sens. Actuators B Chem. 2016,235, 723–731. [CrossRef] 43. Andreadou, M.; Liandris, E.; Gazouli, M.; Taka, S.; Antoniou, M.; Theodoropoulos, G.; Tachtsidis, I.; Goutas, N.; Vlachodimitropoulos, D.; Kasampalidis, I.; et al. A novel non-amplification assay for the detection of Leishmania spp. in clinical samples using gold nanoparticles. J. Microbiol. Methods 2014,96, 56–61. [CrossRef] 44. Anfossi, L.; Di Nardo, F.; Profiti, M.; Nogarol, C.; Cavalera, S.; Baggiani, C.; Giovannoli, C.; Spano, G.; Ferroglio, E.; Mignone, W.; et al. A versatile and sensitive lateral flow immunoassay for the rapid diagnosis of visceral leishmaniasis. Anal. Bioanal. Chem. 2018,410, 4123–4134. [CrossRef] [PubMed] 45. Perinoto, Â.C.; Maki, R.M.; Colhone, M.C.; Santos, F.R.; Migliaccio, V.; Daghastanli, K.R.; Stabeli, R.G.; Ciancaglini, P.; Paulovich, F.V.; De Oliveira, M.C.; et al. Biosensors for efficient diagnosis of leishmaniasis: Innovations in bioanalytics for a neglected disease. Anal. Chem. 2010,82, 9763–9768. [CrossRef] [PubMed] 46. Souto, D.E.; Fonseca, A.M.; Barragan, J.T.; Luz Rde, C.; Andrade, H.M.; Damos, F.S.; Kubota, L.T. SPR analysis of the interaction between a recombinant protein of unknown function in Leishmania infantum immobilised on dendrimers and antibodies of the visceral leishmaniasis: A potential use in immunodiagnosis. Biosens. Bioelectron. 2015,70, 275–281. [CrossRef] [PubMed] 47. Heli, H.; Sattarahmady, N.; Hatam, G.R.; Reisi, F.; Vais, R.D. An electrochemical genosensor for Leishmania major detection based on dual effect of immobilization and electrocatalysis of cobalt-zinc ferrite quantum dots. Talanta 2016 ,156–157, 172–179. [CrossRef] [PubMed] 48. Moradi, M.; Sattarahmady, N.; Hatam, G.; Heli, H. Electrochemical Genosensing of Leishmania major using Gold Hierarchical Nanoleaflets. J. Biol. Today’s World 2016,5, 128–136. [CrossRef] 49. Nazari-Vanani, R.; Sattarahmady, N.; Yadegari, H.; Delshadi, N.; Hatam, G.R.; Heli, H. Electrochemical quantitation of Leishmania infantum based on detection of its kDNA genome and transduction of non-spherical gold nanoparticles. Anal. Chim. Acta 2018 , 1041, 40–49. [CrossRef] 50. Mobed, A.; Mehri, P.; Hasanzadeh, M.; Mokhtarzadeh, A. Binding of Leishmania spp with gold nanoparticles supported polyethylene glycol and its application for the sensitive detection of infectious photogenes in human plasma samples: A novel biosensor. J. Mol. Recognit. 2020,33, e2839. [CrossRef] 51. Diouani, M.F.; Ouerghi, O.; Belgacem, K.; Sayhi, M.; Ionescu, R.; Laouini, D. Casein-Conjugated Gold Nanoparticles for Amperometric Detection of Leishmania infantum. Biosensors 2019,9, 68. [CrossRef] 52. De la Escosura-Muñiz, A.; Baptista-Pires, L.; Serrano, L.; Altet, L.; Francino, O.; Sánchez, A.; Merkoçi, A. Magnetic Bead/Gold Nanoparticle Double-Labeled Primers for Electrochemical Detection of Isothermal Amplified Leishmania DNA. Small 2016 , 12, 205–213. [CrossRef] [PubMed] 53. Moreno, M.; González, V.M.; Rincón, E.; Domingo, A.; Domínguez, E. Aptasensor based on the selective electrodeposition of protein-linked gold nanoparticles on screen-printed electrodes. Analyst 2011,136, 1810–1815. [CrossRef] [PubMed] 54. He, S.; Song, B.; Li, D.; Zhu, C.; Qi, W.; Wen, Y.; Wang, L.; Song, S.; Fang, H.; Fan, C. A Graphene Nanoprobe for Rapid, Sensitive, and Multicolor Fluorescent DNA Analysis. Adv. Funct. Mater. 2010,20, 453–459. [CrossRef] 55. Liu, Z.; Su, X. A novel fluorescent DNA sensor for ultrasensitive detection of Helicobacter pylori. Biosens. Bioelectron. 2017 , 87, 66–72. [CrossRef] [PubMed] 56. Dubertret, B.; Calame, M.; Libchaber, A.J. Single-mismatch detection using gold-quenched fluorescent oligonucleotides. Nat. Biotechnol. 2001,19, 365–370. [CrossRef] [PubMed] 57. Li, H.; Zhang, Y.; Wang, L.; Tian, J.; Sun, X. Nucleic acid detection using carbon nanoparticles as a fluorescent sensing platform. Chem. Commun. 2011,47, 961–963. [CrossRef] [PubMed] 58. Yan, J.-K.; Ma, H.-L.; Cai, P.-F.; Wu, J.-Y. Highly selective and sensitive nucleic acid detection based on polysaccharidefunctionalized silver nanoparticles. Spectrochim. Acta Part A Mol. Biomol. Spectrosc. 2015,134, 17–21. [CrossRef] 59. Bülbül, G.; Hayat, A.; Mustafa, F.; Andreescu, S. DNA assay based on Nanoceria as Fluorescence Quenchers (NanoCeracQ DNA assay). Sci. Rep. 2018,8, 2426. [CrossRef] [PubMed] 60. Loo, A.H.; Sofer, Z.; Bouša, D.; Ulbrich, P.; Bonanni, A.; Pumera, M. Carboxylic Carbon Quantum Dots as a Fluorescent Sensing Platform for DNA Detection. ACS Appl. Mater. Interfaces 2016,8, 1951–1957. [CrossRef] 61. Zhao, H.-Q.; Qiu, G.-H.; Liang, Z.; Li, M.-M.; Sun, B.; Qin, L.; Yang, S.-P.; Chen, W.-H.; Chen, J.-X. A zinc(II)-based two-dimensional MOF for sensitive and selective sensing of HIV-1 ds-DNA sequences. Anal. Chim. Acta 2016,922, 55–63. [CrossRef] 62. Tang, G.; Gao, J.; Wang, C.; Tan, H. Luminescent lanthanide coordination polymer as a platform for DNA colorimetric detection. Sens. Actuators B Chem. 2017,244, 571–576. [CrossRef] Pharmaceutics 2021,13, 491 19 of 19 63. Zhu, C.; Zeng, Z.; Li, H.; Li, F.; Fan, C.; Zhang, H. Single-Layer MoS2-Based Nanoprobes for Homogeneous Detection of Biomolecules. J. Am. Chem. Soc. 2013,135, 5998–6001. [CrossRef] 64. Mo, L.; Li, J.; Liu, Q.; Qiu, L.; Tan, W. Nucleic acid-functionalized transition metal nanosheets for biosensing applications. Biosens. Bioelectron. 2017,89, 201–211. [CrossRef] 65. Zhang, Y.; Liu, S.; Wang, L.; Luo, Y.; Tian, J.; Asiri, A.M.; Al-Youbi, A.O.; Sun, X. Novel Use of Poly(3,4-ethylenedioxythiophene) Nanoparticles for Fluorescent Nucleic Acid Detection. ACS Comb. Sci. 2012,14, 191–196. [CrossRef] [PubMed] 66. Lu, W.; Qin, X.; Chang, G.; Sun, X. 2,4,6-Tris (2-pyridyl)-1,3,5-triazine nanobelts as an effective fluorescent sensing platform for DNA detection. J. Nanosci. Nanotechnol. 2012,12, 2089–2093. [CrossRef] [PubMed] 67. Pedro, G.C.; Gorza, F.D.S.; Da Silva, R.J.; Do Nascimento, K.T.O.; Medina-Llamas, J.C.; Chávez-Guajardo, A.E.; Alcaraz-Espinoza, J.J.; De Melo, C.P. A novel nucleic acid fluorescent sensing platform based on nanostructured films of intrinsically conducting polymers. Anal. Chim. Acta 2019,1047, 214–224. [CrossRef] [PubMed] 68. Andreadou, M.; Liandris, E.; Kasampalidis, I.N.; Taka, S.; Antoniou, M.; Ntais, P.; Vaiopoulou, A.; Theodoropoulos, G.; Gazouli, M.; Ikonomopoulos, J. Evaluation of the performance of selected in-house and commercially available PCR and real-time PCR assays for the detection of Leishmania DNA in canine clinical samples. Exp. Parasitol. 2012,131, 419–424. [CrossRef] [PubMed] 69. Souto, D.E.P.; Silva, J.V.; Martins, H.R.; Reis, A.B.; Luz, R.C.S.; Kubota, L.T.; Damos, F.S. Development of a label-free immunosensor based on surface plasmon resonance technique for the detection of anti-Leishmania infantum antibodies in canine serum. Biosens. Bioelectron. 2013,46, 22–29. [CrossRef] 70. Aldewachi, H.; Chalati, T.; Woodroofe, M.N.; Bricklebank, N.; Sharrack, B.; Gardiner, P. Gold nanoparticle-based colorimetric biosensors. Nanoscale 2018,10, 18–33. [CrossRef] 71. Zhou, W.; Jimmy Huang, P.-J.; Ding, J.; Liu, J. Aptamer-based biosensors for biomedical diagnostics. Analyst 2014 , 139, 2627–2640. [CrossRef] 72. Wang, Y.; Yan, W.; Fu, S.; Hu, S.; Wang, Y.; Xu, J.; Ye, C. Multiple Cross Displacement Amplification Coupled With NanoparticlesBased Lateral Flow Biosensor for Detection of Staphylococcus aureus and Identification of Methicillin-Resistant S. aureus. Front. Microbiol. 2018,9, 907. [CrossRef] 73. Posthuma-Trumpie, G.A.; Korf, J.; Van Amerongen, A. Lateral flow (immuno)assay: Its strengths, weaknesses, opportunities and threats. A literature survey. Anal. Bioanal. Chem. 2009,393, 569–582. [CrossRef] 74. Rivas, L.; Reuterswärd, P.; Rasti, R.; Herrmann, B.; Mårtensson, A.; Alfvén, T.; Gantelius, J.; Andersson-Svahn, H. A vertical flow paper-microarray assay with isothermal DNA amplification for detection of Neisseria meningitidis. Talanta 2018 , 183, 192–200. [CrossRef] 75. Moradi, M.; Sattarahmady, N.; Rahi, A.; Hatam, G.R.; Sorkhabadi, S.M.R.; Heli, H. A label-free, PCR-free and signal-on electrochemical DNA biosensor for Leishmania major based on gold nanoleaves. Talanta 2016 ,161, 48–53. [CrossRef] [PubMed] 76. Bouvier, J.; Schneider, P.; Etges, R.; Bordier, C. Peptide substrate specificity of the membrane-bound metalloprotease of Leishmania. Biochemistry 1990,29, 10113–10119. [CrossRef] 77. Yao, C.; Donelson, J.E.; Wilson, M.E. The major surface protease (MSP or GP63) of Leishmania sp. Biosynthesis, regulation of expression, and function. Mol. Biochem. Parasitol. 2003,132, 1–16. [CrossRef] 78. Bayat, P.; Nosrati, R.; Alibolandi, M.; Rafatpanah, H.; Abnous, K.; Khedri, M.; Ramezani, M. SELEX methods on the road to protein targeting with nucleic acid aptamers. Biochimie 2018,154, 132–155. [CrossRef] [PubMed] 79. Ospina-Villa, J.D.; Zamorano-Carrillo, A.; Castañón-Sánchez, C.A.; Ramírez-Moreno, E.; Marchat, L.A. Aptamers as a promising approach for the control of parasitic diseases. Braz. J. Infect. Dis. 2016,20, 610–618. [CrossRef] [PubMed] 80. Wolter, O.; Mayer, G. Aptamers as Valuable Molecular Tools in Neurosciences. J. Neurosci. 2017 ,37, 2517–2523. [CrossRef] [PubMed] 81. Pan, Q.; Luo, F.; Liu, M.; Zhang, X.-L. Oligonucleotide aptamers: Promising and powerful diagnostic and therapeutic tools for infectious diseases. J. Infect. 2018,77, 83–98. [CrossRef] 82. Ruiz Ciancio, D.; Vargas, M.R.; Thiel, W.H.; Bruno, M.A.; Giangrande, P.H.; Mestre, M.B. Aptamers as Diagnostic Tools in Cancer. Pharmaceuticals 2018,11, 86. [CrossRef] [PubMed] 83. Moreno, M.; Rincón, E.; Piñeiro, D.; Fernández, G.; Domingo, A.; Jiménez-Ruíz, A.; Salinas, M.; González, V.C.M. Selection of aptamers against KMP-11 using colloidal gold during the SELEX process. Biochem. Biophys. Res. Commun. 2003 ,308, 214–218. [CrossRef] 84. Ramos, E.; Piñeiro, D.; Soto, M.; Abanades, D.R.; Martín, M.E.; Salinas, M.; González, V.M. A DNA aptamer population specifically detects Leishmania infantum H2A antigen. Lab. Investig. J. Tech. Methods Pathol. 2007,87, 409–416. [CrossRef] 85. Martín, M.E.; García-Hernández, M.; García-Recio, E.M.; Gómez-Chacón, G.F.; Sánchez-López, M.; González, V.M. DNA aptamers selectively target Leishmania infantum H2A protein. PLoS ONE 2013,8, e078886. [CrossRef] [PubMed] 86. Guerra-Pérez, N.; Ramos, E.; García-Hernández, M.; Pinto, C.; Soto, M.; Martín, M.E.; González, V.M. Molecular and Functional Characterization of ssDNA Aptamers that Specifically Bind Leishmania infantum PABP. PLoS ONE 2015 , 10, e0140048. [CrossRef] [PubMed] 87. Frezza, V.; Pinto-Díez, C.; Fernández, G.; Soto, M.; Martín, M.E.; García-Sacristán, A.; González, V.M. DNA aptamers targeting Leishmania infantum H3 protein as potential diagnostic tools. Anal. Chim. Acta 2020,1107, 155–163. [CrossRef] [PubMed]