Leishmaniasis is a group of diseases caused by parasites from over 20 Leishmania species, which present two main clinical forms: cutaneous leishmaniasis (affecting the skin and/or mucosa) (CL) and visceral leishmaniasis (VL), affecting the lymphohematopoietic system. Leishmaniasis represents a significant global health challenge, primarily affecting the Americas, East Africa, North Africa, and West and Southeast Asia (SAÚDE OO-OP-AD, 2021), and is endemic in 90 countries.

CL is a non-contagious infectious disease that causes ulcers on the skin and/or mucosa and is transmitted to humans through the bite of infected female sandflies of the genus Lutzomyia, commonly known as sandflies (Silva et al., 2021). In Brazil, the main Leishmania species responsible for infection are Leishmania (Leishmania) amazonensis, L. (Viannia) guyanensis, and L. (V.) braziliensis. However, other species also occur, such as L. naiffi, L. lainsoni, L. lindenbergi, and L. shawi, which are also found in Brazil. WHO currently estimates that more than 1 billion people live in endemic areas and are at risk of infection, with approximately 30,000 new cases of VL and 1 million new cases of CL occurring annually (World Health Organization, 2025a). In Brazil, an average of approximately 21,000 cases is reported annually, with the North region exhibiting the highest incidence rate (46.4 cases per 100,000 inhabitants), followed by the Center-West (17.2 cases per 100,000 inhabitants) and Northeast (eight cases per 100,000 inhabitants) (Ministério da Saúde, 2025b).

In the Americas, VL is characterized as a chronic zoonotic disease that can involve systemic manifestations and may be fatal in up to 90% of cases if not properly treated. VL is also transmitted to humans through the bite of infected female sandflies of the genus Lutzomyia, with Lutzomyia longipalpis being the primary species involved. Transmission occurs when the female sandfly feeds on infected dogs (urban reservoirs) or wild animals (sylvatic reservoirs) and subsequently bites a human (Ministério da Saúde, n.d.(c)). It is estimated that 50,000 to 90,000 new cases occur worldwide each year, with the majority in East Africa, India, and Brazil (World Health Organization, 2023a). In Brazil, an average of approximately 2.000 new cases is reported annually, particularly in the North and Northeast regions, caused by Leishmania infantum (Marcondes and Day, 2019; Lopez et al., 2025).

The transmission cycles of CL and VL share similar characteristics, involving a mammalian host and a vector. During the blood meal, the sandfly injects infective promastigote forms into a susceptible mammal. These promastigotes are phagocytosed by macrophages and other phagocytic cells, transforming into amastigote forms that multiply by binary fission within a parasitophorous vacuole until causing cell lysis and subsequent phagocytosis by new cells, leading to eventual parasitemia (Esch and Petersen, 2013).

Infection of sandflies occurs through a blood meal taken from a host, ingesting infected macrophages. Once inside the insect vector, the amastigotes transform into promastigotes in the midgut, reproduce, and subsequently migrate to the valve stomodeal, where they develop into infective metacyclic promastigotes until the next blood meal (Esch and Petersen, 2013). It is noteworthy that domestic dogs are considered the primary urban reservoir of VL, whereas foxes and marsupials serve as sylvatic reservoirs, and generally, cases in dogs precede human cases (Costa et al., 2018; Brasil MdS, 2014; Camargo and Langoni, 2006).

The symptoms of CL are lesions on the skin and/or mucosa, which may be single, multiple, disseminated, or diffuse, typically presenting as painless ulcers (Ministério da Saúde, n.d.(b)). In contrast, VL is characterized as a systemic infection, causing prolonged fever, hepatosplenomegaly, weight loss, weakness, muscle fatigue, anemia, leukopenia, thrombocytopenia, hemorrhagic phenomena, among other clinical signs (Ministério da Saúde, n.d.(c)).

In humans, the pharmacological treatment of visceral and cutaneous leishmaniasis includes pentavalent antimonials (Pentostam^® and Glucantime^®), amphotericin B, paromomycin, and miltefosine (Santos et al., 2023). In Brazil, miltefosine is the only drug administered orally and is currently also used for the treatment of canine visceral leishmaniasis, reducing the parasite load in the skin and bone marrow and it can also be combined with allopurinol to enhance the therapeutic response (Vaz et al., 2023). However, these treatments have limitations, including high cost, parasite resistance, therapeutic failure and severe adverse effects (Santos et al., 2023).

Accordingly, new orally administered treatment regimens are being investigated, including compounds such as 2,4,5-trisubstituted benzamides, 4,7,9-trisubstituted benzoxazepines, and farnesol (Kadayat et al., 2024; Kim et al., 2023; Sharma et al., 2023).

The 2,4,5-trisubstituted benzamides are benzamide derivatives that exhibit in vitro activity against L. mexicana amastigotes (EC₅₀ = 0.66 μM) and display high oral bioavailability (80%) (Kim et al., 2023; Sharma et al., 2023). 4,7,9-trisubstituted benzoxazepines, derived from 4-[(3,5-dimethyl-4-isoxazolyl)acetyl]-9-[(1-methyl-3-piperidinyl)methoxy]-7-(5-methyl-2-thienyl)-2,3,4,5-tetrahydro-1,4-benzoxazepine, showed an EC₅₀ of 2,3 μM against L. mexicana amastigotes and demonstrated good aqueous solubility (Kadayat et al., 2024). Farnesol, a sesquiterpene found in plants such as citronella, cyclamen, balsam, and musk, as well as in various essential oils, and known for its low toxicity, exhibited inhibitory dose-dependent activity against L. major promastigotes (EC₅₀ = 175.7 μM) and amastigotes (EC₅₀ = 945 μM), with a selectivity index (SI = 5.65). Its mechanism of action includes induction of apoptosis and inhibition of the enzyme lanosterol 14-demethylase, ultimately disrupting ergosterol biosynthesis (Kim et al., 2023; Sharma et al., 2023).

Vaccine development is challenging due to the complex life cycle of the parasite. The Sucrose Non-Fermenting 1 (SNF1) kinase protein has emerged as a potential target for vaccine development. Deletion of this gene using the CRISPR-Cas9 technique resulted in reduced promastigote growth, a lower proportion of metacyclic forms, and morphological deformation of mitochondrial cristae. In mouse models, this deletion also led to a reduction in lesion size (Shoeran et al., 2025).

Diagnosis of CL is performed using direct parasitological methods, such as lesion edge scrapings or imprint biopsy techniques. Histopathological examination, parasite isolation in culture, serology, and PCR can also be used (Xavier et al., 2006; Saldanha et al., 2017). VL diagnosis is also conducted via parasitological methods through the visualization of amastigote forms in collected material. According to the Brazilian Ministry of Health, bone marrow aspirate is considered the biological material of reference for direct parasitological diagnosis. However, other samples can be used to visualize Leishmania amastigotes, such as lymph nodes and spleen, which require procedures to be performed in a hospital setting and under surgical conditions. Other tests for the diagnosis of LV include: serology, in addition to PCR or rapid immunochromatographic tests (Gao et al., 2015; Sakkas et al., 2016).

Both forms of leishmaniasis can be controlled and prevented through vector management, including practices such as cleaning yards and vacant lots, removing organic material, using insecticides and repellents in areas with high case numbers, and implementing surveillance and environmental education measures in affected regions (World Health Organization, n.d.(c)).

Chagas Disease, also known as American trypanosomiasis, is an infection caused by the protozoan Trypanosoma cruzi. It presents an acute phase, which may be symptomatic or asymptomatic, and a chronic phase, which can be asymptomatic or manifest with cardiac and/or digestive clinical symptoms (Ministério da Saúde, n.d.(a)). WHO estimates that approximately 7 million people worldwide are infected with T. cruzi, with around 10,000 deaths reported in 2017. Additionally, the disease has been detected in 44 countries (World Health Organization, n.d.(a)). In Brazil, during the period from 2023 to 2024, 5.460 cases of chronic Chagas disease were reported across 710 municipalities, with most cases occurring in urban areas (Ministério da Saúde, 2024a).

The transmission cycle of Chagas disease begins with the vector, an infected triatomine bug, during a blood meal on a host. The insect vector releases trypomastigote metacyclic forms in the feces near the bite site, which causes a minor injury, allowing the trypomastigotes to enter the host and invade nearby cells, differentiating into intracellular amastigotes. These amastigotes multiply by binary fission and subsequently differentiate into trypomastigotes, which are released into the bloodstream to infect new cells, transforming back into intracellular amastigotes. The triatomine becomes infected when feeding on a host with parasitemia (Prevention CfDCa, 2021).

It is noteworthy that Chagas disease can be transmitted through several routes, the main ones being: vector-borne, via direct contact with triatomine feces; oral, through ingestion of food contaminated with T. cruzi; vertical, via transmission from mother to baby during pregnancy or childbirth; or through blood transfusion, organ transplantation, and, less commonly, accidental contact of wounds or mucosa with contaminated material (Ministério da Saúde, n.d.(a)).

The most common symptoms of Chagas disease during the acute phase include prolonged fever lasting more than seven days, headache, severe weakness, swelling of the face and legs, and the appearance of a chagoma, the classical lesion of the disease, which resembles a boil at the site of the triatomine bite. Another characteristic symptom of the acute phase is Romaña's sign, a classic indicator of acute Chagas disease. After the acute phase, if the individual does not receive timely and appropriate treatment, the chronic phase of the disease may develop, initially remaining asymptomatic. However, over the years, infected individuals may develop cardiac, digestive, or cardio digestive complications (Ministério da Saúde, n.d.(a)).

Diagnosis of Chagas disease is based on the presence of suggestive signs and symptoms and an epidemiological history compatible with exposure to outbreaks. In the acute phase, diagnosis is generally performed through direct parasitological methods such as thick blood smear and peripheral blood smear and/or detection of anti-T. cruzi IgM antibodies. In the chronic phase, diagnosis is performed through the detection of antibodies using indirect immunofluorescence, hemagglutination, and ELISA assays, as well as DNA detection by PCR. The establishment of cell culture (blood culture) can also be employed (Souza and Amato Neto, 2012; Ramírez et al., 2009; Duarte et al., 2014).

In Brazil, for the treatment of Chagas disease, the first-line therapy is benznidazole, administered in two doses per day. As a second-line therapy, nifurtimox is recommended, administered in three doses per day for up to 60 days (Altcheh et al., 2025; Freire et al., 2025; Díaz-Menéndez et al., 2025). These drugs are capable of reducing parasitemia, and when cure is achieved, seroconversion occurs, with antibody levels becoming seronegative (Altcheh et al., 2025).

Benznidazole is effective in the acute phase of the disease; however, prolonged treatment allows the elimination of the bloodstream forms and reduces the tissue form of the parasite (Freire et al., 2025). This drug presents, as a disadvantage in the chronic phase, a reduced ability to eliminate amastigote forms, and it also has several side effects, such as neurological, dermatological, and gastrointestinal alterations, sometimes requiring treatment discontinuation. However, new drugs and therapeutic combinations have been studied (Díaz-Menéndez et al., 2025). The combination of benznidazole with curcumin showed a cure rate of 83.3%, while amiodarone and atorvastatin resulted in reduced parasitemia in the blood and demonstrated important cardioprotective effects in heart failure and arrhythmias (Freire et al., 2025).

Malaria is an infectious disease caused by parasites of the genus Plasmodium and is transmitted to humans through the bite of infected female mosquitoes of the genus Anopheles, which are most active at dusk and dawn (Ministério da Saúde, n.d.(d)). In Brazil, the most important Plasmodium species are P. malariae, P. vivax, and P. falciparum, while P. ovale can occasionally be diagnosed in the country but is more frequently found in Africa (Oliveira-Ferreira et al., 2010; Limongi et al., 2014; da Silva, 2010). Among these species, P. falciparum is considered the most lethal and is the predominant species in Africa, while P. vivax predominates in most countries outside sub-Saharan Africa (Prugnolle et al., 2011; Howes et al., 2016).

According to WHO, an estimated 263 million malaria cases occurred worldwide in 2023, resulting in approximately 597,000 deaths, with Africa reporting 94% of the cases. In Brazil, the Amazon region is considered endemic, accounting for about 99% of cases. In 2023, more than 140,000 autochthonous cases (those acquired in the same location where they were diagnosed) were reported, along with 63 deaths, of which 17.3% were caused by P. falciparum and 82.7% by P. vivax and other species (Ministério da Saúde, n.d.(e)).

The malaria transmission cycle involves two hosts. During a blood meal, an infected female Anopheles mosquito inoculates sporozoite forms into the host. These parasites infect liver cells and transform into schizonts, which rupture and release merozoite forms. After the initial replication in the liver, the parasites undergo asexual multiplication in erythrocytes, where merozoites infect red blood cells. Ring-stage trophozoites then mature into schizonts, which rupture and release merozoites back into the bloodstream to infect new red blood cells. It is this rupture that leads to the clinical manifestations of malaria. Some parasites differentiate into male (microgametocytes) and female (macrogametocytes) gametocytes, which are ingested by the Anopheles mosquito during a blood meal from an individual with parasitemia. In the mosquito, parasite multiplication occurs through the sporogonic cycle, in which male gametes penetrate female gametes to form zygotes. These zygotes become motile and elongated, known as ookinetes, which invade the mosquito gut wall and develop into oocysts. The oocysts grow, rupture, and release sporozoites, which migrate to the mosquito's salivary glands and are inoculated into a new host (Tuteja, 2007; García-Basteiro et al., 2012).

The most common symptoms of malaria are high fever, chills, shivering, sweating, and headache, which may occur cyclically. Less common symptoms include nausea, vomiting, fatigue, and loss of appetite. Severe malaria is characterized by one or more of the following symptoms: prostration, convulsions, altered consciousness, hypotension, dyspnea or hyperventilation, and hemorrhages (Ministério da Saúde, n.d.(d)). It is noteworthy that individuals who have experienced multiple malaria episodes or are continuously exposed may develop naturally acquired immunity, providing protection against clinical symptoms and resulting in low parasitemia (Doolan et al., 2009).

Malaria diagnosis is performed through the visualization of the parasite in parasitological examinations, with the thick blood smear method considered the gold standard, although it can also be assessed using peripheral blood smears (Makler et al., 1998; O'Meara et al., 2006). Rapid tests detecting Plasmodium antigens are currently available, and in some cases, PCR may also be performed (Opoku Afriyie et al., 2023; de Fátima Ferreira-da-Cruz et al., 2025; Andrade et al., 2010).

Therapeutic strategies for malaria involve the use of artemisinin-based drugs or combination therapies, such as (1) artesunate and amodiaquine, (2) artemether and lumefantrine, (3) dihydroartemisinin and piperaquine, (4) artesunate and sulfadoxine–pyrimethamine, (5) artesunate and pyronaridine (Okombo and Fidock, 2025). The therapeutic combination reduces the risk of treatment failure, enhances parasite clearance, and improves treatment tolerance (Sudhakaran, 2025). In addition to therapeutic combinations, other drugs have been identified as promising candidates, including cabamiquine (a quinoline-carboxamide), cipargamin (a spiroindolone), and ganaplacide (an imidazolopiperazine) (Okombo and Fidock, 2025).

Moreover, vaccine development strategies are being explored as preventive measures (Naghizadeh et al., 2025). The transmission-blocking malaria vaccine (ProC6C-AlOH/Matrix-M™) was developed by incorporating three parasite proteins (Pfs230-Pro, Pfs48/45-6C, and CSP) and formulating them with two adjuvant combinations: aluminum hydroxide alone (ProC6C-AlOH) and aluminum hydroxide combined with the saponin-based Matrix-M™ adjuvant (ProC6C-AlOH/Matrix-M™) (Naghizadeh et al., 2025). Currently in Phase 1 clinical trials, this vaccine aims to prevent infection across multiple stages of the parasite's life cycle and has been shown to be safe, with the potential to induce high levels of specific IgG (Naghizadeh et al., 2025). Therefore, the development of new therapeutic and vaccine strategies is essential for controlling and reducing the transmission of the disease.

ELISA is an immunological technique widely used to detect the presence of proteins, antibodies, hormones, and other molecules in a sample. It is based on the highly specific and sensitive interaction between antigens and antibodies, mediated by enzymatic reactions (Pereira Filho, 2023). The assay involves the incubation of reagents, and its final outcome is the development of a measurable color change, which can be quantified using a spectrophotometer. This quantification enables accurate interpretation of the results. As a diagnostic method, ELISA aims to assess specific concentrations of antigens or antibodies in order to determine whether the analyzed serum can show the infection and to evaluate the level of immune protection (Crowther, 2008). Moreover, due to its high reproducibility and ability to generate a large number of results within a single experiment, this technique is widely employed in both scientific research laboratories and clinical diagnostic settings (Hosseini et al., 2017).

The ELISA assay can be either qualitative or quantitative, and in general, the assay can be categorized into four main formats: direct ELISA, indirect ELISA, sandwich ELISA, and competitive ELISA (Figure 3). Direct ELISA is the most practical form of the assay and aims to verify the presence or even quantify the amount of antibody, allowing the identification of whether an individual or animal has been exposed to a given pathogen. In this assay, antigens derived from the sample of interest are immobilized, followed by the addition of a primary antibody conjugated to an enzyme (peroxidase or alkaline phosphatase), which enables the reaction and color change upon contact with the substrate due to enzymatic catalysis. The intensity of the color is visually estimated and is proportional to the concentration of the antibody under investigation. Since only one antibody is added, there is no possibility of cross-reactivity in this type of assay. Furthermore, it can be performed in a shorter time compared to other formats (Lin, 2015). On the other hand, the assay has disadvantages such as low sensitivity, due to the use of only one antibody, and consequently reduced signal amplification (Crowther, 2008; Gan and Patel, 2013).

The indirect ELISA involves the addition of the antigen for plate sensitization, followed by the addition of the sample that may contain the antibody of interest. Afterwards, a second antibody (secondary antibody conjugated to an enzyme), which is specific to the primary antibody, is added. Finally, the substrate is introduced, and the enzyme generates the detection signal resulting from the chemical reaction: enzyme, hydrogen peroxide, and substrate. The advantages of Indirect ELISA include versatility, as a single primary antibody can be used to detect different antigens; and sensitivity, as it is generally more sensitive than direct ELISA by allowing detection of multiple secondary antibodies bound to a single primary antibody, amplifying the detection signal (Lin, 2015; Tabatabaei and Ahmed, 2022). However, signal amplification may also increase the likelihood of cross-reactivity, thereby raising background signal levels (Crowther, 2008).

The sandwich ELISA is widely used to detect specific antigens at low concentrations. In this method, two different specific antibodies are employed: a capture antibody and a detection antibody. The capture antibody is immobilized on the plate, after which the sample is added. The antigen present binds to the capture antibody, followed by the addition of the detection antibody conjugated to an enzyme, thus forming an antibody-antigen–antibody complex. After washing, the substrate is added, and the enzyme produces a colorimetric detection signal. Sandwich ELISA has advantages such as high specificity, since two different antibodies bind to distinct epitopes of the antigen, enabling more specific detection and minimizing interference and cross-reactivity. It is often considered more sensitive than Direct or Indirect ELISA (Pereira Filho, 2023). The disadvantages include the requirement for two specific antibodies that must not compete for the same epitope, which can be challenging and costly, as well as the limitation that this assay cannot be applied to antigens with only one epitope, since they do not allow simultaneous binding of two antibodies (Crowther, 2008; Gan and Patel, 2013).

In the competitive ELISA, the antigen is first incubated with a specific antibody conjugated to an enzyme. This antigen–antibody complex is then added to a plate pre-coated with a capture antibody. The labeled antigen competes with the antigen from the sample for binding to the capture antibody on the plate surface. The higher the amount of antigen in the sample, the less labeled antigen will bind to the capture antibody. In this case, the enzymatic activity is measured as inversely proportional to the amount of antigen present in the sample. Although Competitive ELISA is less common than other formats, it presents specific advantages such as broad applicability, being suitable for the detection of small, poorly immunogenic antigens where other ELISA formats may not be appropriate, and high specificity due to the competition principle (it can be highly sensitive for detecting specific antigens) (Pereira Filho, 2023; Lequin, 2005). However, if there are substances in the sample capable of nonspecific competition, this may compromise the success of the technique. In addition, it is considered less sensitive than Sandwich ELISA, precisely due to the competition process, which naturally results in a lower signal (Lequin, 2005).

The selection of the most appropriate format depends directly on the specific experimental question being addressed. Each variation offers the possibility of optimizing aspects of the test, such as enhancing sensitivity, diversifying the types of antibodies employed, and amplifying the detection signal. Consequently, ELISA stands out as a highly versatile technique, applicable across a wide range of research and diagnostic contexts (Pereira Filho, 2023).

The execution of the ELISA test follows a sequence of essential steps. Initially, sensitization is performed, during which the antigen or antibody is adsorbed onto the surface of the plate. The analysis is typically conducted in a microtiter plate specifically coated for the assay of interest, where specific antigens or antibodies are immobilized on the well surfaces, creating a “coating” capable of capturing the target. These plates are usually made of polystyrene, a material with a protein-adsorbing surface, and feature either flat- or U-shaped bottoms (Engvall et al., 1971). The type of ELISA being performed determines which antigens or antibodies are required at this stage, as well as their concentrations and amounts.

The next step involves blocking the free binding sites to prevent nonspecific interactions. For this purpose, bovine serum albumin (BSA) is commonly used, as it adheres to the plate and prevents unwanted binding of reagents that could otherwise generate false results (Lequin, 2005). After blocking, the plate is washed with PBS containing Tween 20 to remove excess blocking agent. Between each step, careful washing is required to eliminate unbound reagents from the supernatant, preventing interference with the assay outcome. Washing is performed with a buffer solution supplemented with a nonionic detergent, typically phosphate-buffered saline (PBS), prepared by diluting a 10 × stock solution to 1 × , in combination with Tween 20 (Crowther, 2008). The number of washes depends on the specific protocol being followed, and they may be performed using an automated plate washer or manually with pipettes and other tools.

Subsequently, the samples can be incubated, followed by another wash with PBS + Tween 20 to remove unbound material. A secondary antibody conjugated to an enzyme is then added, followed by additional washes to eliminate unbound antibodies. To generate a colorimetric signal at the end of the experiment, the primary antibody (in direct ELISA) or the secondary antibody (in indirect, sandwich, or competitive ELISA) is conjugated to an enzyme, most commonly horseradish peroxidase (HRP) or alkaline phosphatase (AP) (Crowther, 2008).

Finally, a chromogenic substrate is added, which reacts with the enzyme to produce a colorimetric signal. Depending on the enzyme–substrate combination, the final product may exhibit a blue or orange color (Pereira Filho, 2023). The most frequently used substrates are o-phenylenediamine dihydrochloride (OPD) and 3,3′,5,5′-tetramethylbenzidine (TMB) (Crowther, 2008). To allow proper measurement, the enzymatic reaction must be stopped before the plate is read in an ELISA plate reader. This is achieved by adding an acid, most commonly sulfuric acid (H₂SO₄) (Crowther, 2008) or hydrochloric acid (HCl) (Gan and Patel, 2013). After stopping the reaction, the plate must be read in an ELISA reader within 30 min. The wavelength selected for measurement depends on the stop solution employed in the assay.

In pathogen diagnosis, ELISA is an indispensable technique, being employed from the confirmation of protozoan presence to the quantification of its peptides, particularly in relation to the most common protozoal infections in Brazil. Numerous examples could be cited regarding the role of ELISA in diagnosing these diseases. In leishmaniasis, ELISA can be applied in patient screening (Bia et al., 2025), in diagnostic confirmation (Casas et al., 2024), after chromatography (Pradella et al., 2025), and in the quantification of molecules such as cytokines following disease resolution (Jawalagatti et al., 2023). In Chagas disease, ELISA can be used in a variety of ways, from the investigation of antigens or antibodies (Suescún-Carrero et al., 2022) to direct disease diagnosis, being the most widely employed serological technique (Ossowski et al., 2024). In malaria, ELISA plays multiple roles, from the detection of antigens in the human host or in the vector, to the identification of specific antibodies in the human host (Hendershot et al., 2021; Jang et al., 2022), and even in epidemiological studies to assess population exposure to the parasite (Ventocilla et al., 2024).

Over the years, the continuous evolution and refinement of this technique has broadened its range of applications, enhancing its sensitivity, specificity, and reliability, assisting in improving the technique in order to avoid false positives or negatives. Despite some limitations, the benefits associated with the ELISA assay, such as ease of implementation, high reproducibility, and relatively low cost, significantly outweigh its disadvantages. For these reasons, ELISA remains one of the most important analytical tools employed in clinical and research laboratories worldwide (Pereira Filho, 2023).

ELISpot assay is an immunoassay used to quantify analyte-secreting cells and these analytes may include cytokines, immunoglobulins, or other target proteins that are secreted upon specific cellular stimulation and subsequently captured by specific antibodies (Gertow, 2022). Since this technique employs enzymes conjugated to antibodies and results in the formation of precipitates in the form of small dots or “spots,” researchers named the assay ELISPOT, from the English enzyme-linked immunospot (Czerkinsky et al., 1983). Since then, the method has been refined to reach its current format, available as commercial kits or with separate reagents, in which, similar to the ELISA technique, consists of an immunochemical “sandwich” combining antigens with pairs of antibodies (capture and detection), as illustrated in Figure 4.

Despite this similarity, it is important to emphasize that these assays provide different types of information and can be used in a complementary manner: considering a given cytokine, for example, ELISA enables the measurement of the actual concentration of the molecule present in the supernatant, i.e., the total amount of cytokine secreted. In contrast, ELISPOT allows the determination of the frequency of the cells secreting that particular cytokine (Kalyuzhny, 2005). Because ELISPOT can detect the capacity of even a single cell to secrete cytokines, it is characterized as a much more sensitive technique—up to 100 times more sensitive than an ELISA assay—thereby enabling the detection of cytokines at levels previously undetectable in the supernatant (GEGINAT GpS, 2005).

In general, the ELISPOT assay can be divided into different stages that take up to 3 days. Briefly, it begins with the coating phase, in which the plate is sensitized with the capture antibody. For this, within a biological safety cabinet, 96-well ELISPOT plates containing a polyvinylidene fluoride (PVDF) membrane are pretreated with ethanol, followed by the addition of the capture antibody and subsequent overnight incubation at 4–8 °C. This sensitization step is unnecessary when using pre-coated ELISPOT plates, a more expensive material, that allows one less day of experiment. On the second day the selected stimulus (e.g., synthetic peptides corresponding to epitopes of interest) is added first, followed by the cells (typically peripheral blood mononuclear cells, PBMCs). In a CO₂ incubator (5% CO₂, 37 °C), the stimulated cells secrete the analyte. This secretion period may range from 12 to 48 h, depending on the analyte of interest. On the third day, after cell removal, the detection antibody specific to the analyte of interest is added, followed by an enzyme conjugate and a chromogenic substrate for the enzyme, BCIP/NBT-plus. Upon reaction, this substrate is converted into a purple precipitate that deposits onto the PVDF membrane in the form of spots and the membrane must be washed with water and allowed to dry completely for at least 24 h before reading the plate in an ELISPOT reader. After drying, the plates can be stored for long periods, thus allowing the technique to be directly performed in endemic areas and the plates can be transferred for analysis at research centers (Ji and Forsthuber, 2016) (Figure 5).

Several studies in this field have shown that the frequency of IFN-γ-producing CD8+ T cells specific for T. cruzi is inversely correlated with the severity of chronic Chagas disease (Laucella et al., 2004). Similarly, research comparing children infected with T. cruzi at early stages to adults in the chronic phase demonstrated that the cellular immune response in children is characterized by polyfunctional T cells, with IFN-γ production up to five times higher than in adults (Albareda et al., 2013). ELISPOT assays have also shown that patients with high pre-treatment frequencies of IFN-γ- and IL-2-producing T cells who received benznidazole exhibited progressively lower antibody titers after treatment completion, as assessed by serology, the current standard for indicating cure through the absence of specific antibodies (Alvarez et al., 2016). These findings, together with other techniques, can contribute to a more precise diagnosis of Chagas disease. There are also studies reporting the use of the ELISPOT technique for the investigation of leishmaniasis, as in the study of the follow-up of two pregnant women with cutaneous leishmaniasis, in which ELISPOT assays were used. In this paper, it was observed that IFN-γ levels increased as lesions healed. This indicates alterations in the maternal immune system, affect susceptibility to Leishmania braziliensis and result in the worsening of the disease (Conceição-Silva et al., 2013). Although the technique was not used for diagnosis itself, it was associated with other techniques in the context of prognosis, a process closely linked to diagnosis and essential for the determination of disease progression. From the perspective of malaria, to date, studies described in the scientific literature focus mainly on aspects such as pathogenesis, epitope identification, and vaccine development (Ventocilla et al., 2024; Atre et al., 2019; John et al., 2004; Ganeshan et al., 2016). Despite the limited number of studies using ELISPOT as an ally in diagnosis, based on the studies cited, this technique may support broader applications, including disease monitoring, assessment of treatment efficacy, and determination of patient cure, as already demonstrated in various tropical diseases (Lima-Junior et al., 2017).

Direct parasitological tests, including histopathology, immunohistochemistry, cytology, and immunocytochemistry, are considered the gold standard for the diagnosis of parasitic diseases. Similar to histopathology, immunohistochemistry requires tissue collected by biopsy, necessitating trained professionals to perform the procedure (Conceição-Silva, 2024). Thus, the Immunohistochemistry (IHC) is a technique that employs the antigen–antibody reaction to detect target molecules in tissue sections. It allows the in situ identification of various structures that may be associated with different diseases and is widely used in anatomical pathology (Ferro, 2014). In the context of diagnosing parasitic diseases, IHC is used to detect antigens of the specific parasite targeted by the test, offering greater specificity compared to conventional histopathology and allowing the differentiation of morphologically similar parasites (Ezyaguirre et al., 2011; Conceição-Silva, 2024). It is very useful for identifying parasites in low amounts that may be difficult to detect using routine staining techniques (Ezyaguirre et al., 2011). The IHC technique can be performed using different methods to identify a specific antigen. However, IHC is considered a complex technique, and several factors can affect the quality of staining. These factors range from early steps, such as tissue fixation and antigen retrieval, to later steps, such as the choice of detection method (Nielsen, 2021). In the direct method, the primary antibody conjugated to a label bind directly to the target antigen, eliminating the need for a secondary antibody. In the indirect method, a primary antibody detects the target antigen, followed by a secondary antibody raised against the immunoglobulin of the species in which the primary antibody was produced. The secondary antibody must be conjugated to a substance that enables visualization of the antigen–antibody complex (Ferro, 2014). In addition to direct and indirect methods, other methods can also be employed, including: Peroxidase Anti-Peroxidase (PAP) method, Alkaline Phosphatase Anti-Alkaline Phosphatase (APAAP) method, avidin-biotin methods, and polymer methods (Murphy et al., 1997).

In the initial steps of the technique, the sample preparation and tissue fixation, are crucial for the preservation, stabilization, and protection of samples to be used in subsequent analyses (Ferro, 2014). Two types of fixations can be employed for IHC: the tissue can be cryopreserved at low temperatures, embedded in an appropriate resin, or fixed in formaldehyde and subsequently embedded in paraffin (Conceição-Silva, 2024). Cryopreservation offers the advantage of a faster IHC procedure compared to paraffin-embedded material. However, the freezing process can cause tissue damage, affecting tissue integrity and, consequently, IHC performance (Colley and Ronald, 2021). The most commonly used fixative is 10% neutral-buffered formalin, which penetrates tissues effectively and preserves morphological details for subsequent paraffin embedding. However, antigens can be masked by formaldehyde, which causes conformational changes in epitopes, resulting in the loss of the antibody's ability to interact with the target antigen, requiring antigen retrieval (Shi and Taylor, 2021).

The antigen retrieval involves treating the tissue to restore the structure of the protein that has been masked by formaldehyde (Ferro, 2014). This can be achieved through enzymatic digestion or heat-induced methods (Shi and Taylor, 2021).

This process can reverse structural modifications induced by formalin, provided that the primary protein structure, determined by the amino acid sequence, remains intact (Ferro, 2014). Optimal conditions for antigen retrieval depend on the antibody to be used. Therefore, when selecting an antibody, it is essential to verify its suitability for paraffin-embedded material and determine the most appropriate retrieval method.

The antibody selection depends on the laboratory, considering factors from tissue processing and fixation to tissue species. Correct dilutions prepared consistently ensure high specificity with minimal background. The dilution, incubation time, and temperature of the primary antibody are also crucial steps. There is an inverse relationship between antibody concentration and incubation time: higher concentrations require shorter incubation (Jensen, 2021). Regarding detection methods, in the case of an antibody conjugated to an enzyme (HRP), a chromogen (DAB or AEC) must be added for the development of the enzymatic reaction. The oxidation by Peroxide (HRP) results in a brown (DAB) or red (AEC) precipitate, indicating the parasite localization in the tissue (Jensen, 2021). Finally, many counterstains can be used in IHC, with hematoxylin being the most common. Nuclear staining in blue improves visualization of tissue morphology and provides contrast with the brown DAB or red AEC staining (Jacobsen et al., 2021). The Figure 6 provides a summary representation of the IHC procedure.

In Leishmania infection, immunohistochemistry is considered a reference test for parasitological diagnosis, allowing correlation of the parasite with tissue lesions and the detection of active infection (Conceição-Silva, 2024). In the case of chronic granulomatous leishmaniasis with a small number of parasites in the sample, immunohistochemical staining has been a very useful diagnostic technique for parasite identification (Ezyaguirre et al., 2011). However, although it is a more sensitive technique in relation to histopathology, there may be cross-reactions with other pathogens such as Trypanosoma cruzi, requiring additional tests for an accurate diagnosis (Ferreira et al., 2022). These cross-reactions may be caused by the use of polyclonal antibodies or hyperimmune sera processed in-house, which can produce nonspecific bindings, considered one of the limitations of the technique (Conceição-Silva, 2024).

Flow cytometry is a methodology capable of simultaneously measuring, within a short period of time, multiple physical and biological characteristics of a single cell or particle, such as size, granularity, morphology, and function, as the cell flows in suspension (Adan et al., 2017). This measurement is achieved through the detection of fluorescence emission from fluorescent molecules—known as fluorochromes—in samples previously prepared and labeled with monoclonal antibodies conjugated to fluorochromes of interest (Ferraz and Bertho, 2013). Due to its ability to analyze several parameters simultaneously at the single-cell level, flow cytometry has become an essential technique for multiparametric analyses, and is widely employed in the immunophenotyping of peripheral blood cells, apoptosis assays, hematology, immunology, cytokine detection, immunogenetics, among others (dos Santos et al., 2017).

The basic principle of flow cytometry relies on the passage of labeled cells in suspension through a laser beam that excites the fluorochrome(s), which in turn emit light at specific wavelengths (colors) captured by strategically positioned detectors. The fluorochromes are typically conjugated to monoclonal antibodies used to label cells, with each fluorochrome exhibiting a distinct excitation and emission spectrum (Flores-Montero et al., 2019; Bertho et al., 2024; Actor, 2024). These conjugated antibodies specifically interact with molecular targets located on the cell surface or within the cytoplasm. Such targets are known as clusters of differentiation (CDs) (Bertho et al., 2024). Thus, combining multiple anti-CD antibodies conjugated to different fluorochromes enables the simultaneous identification of diverse cell populations and their functional states within a single sample in a relatively short time frame.

The detectors, photomultiplier tubes (PMTs), measure the light absorbed or scattered by the sample and convert it into electronic signals that are processed by a computer, enabling morphological and functional analyses using dedicated multiparametric software. The results are commonly displayed as fluorescence intensity plots, depending on the fluorochrome(s) employed, providing information on cellular morphology such as size and granularity (Bajgelman, 2019).

An essential step in flow cytometry is the preparation of biological samples, such as peripheral blood, cell cultures, cerebrospinal fluid, tissue, and bone marrow, among others. To allow proper evaluation, cell suspensions are required; therefore, sample dissociation is necessary to prevent the formation of cellular aggregates that may damage the instrument and generate unreliable results (Ferraz and Bertho, 2013). Additional procedures may be required during sample preparation, including the use of trypsin to detach adherent cells, removal of unwanted contaminating material, or lysis of red blood cells in peripheral blood (Bio Rad Antibodies, n.d.). Subsequently, molecular targets to be labeled with monoclonal antibodies must be selected. The choice of fluorochromes depends on the optical configuration of the flow cytometer (Ferraz and Bertho, 2013).

One of the most commonly applied protocols in flow cytometry is the labeling of cell surface proteins, as the plasma membrane is readily accessible to antibodies (Adan et al., 2017). Intracellular proteins can also be detected by flow cytometry; however, this requires that cells be treated with protein transport inhibitors to ensure retention of secreted proteins. The cells are then fixed, typically with formaldehyde, and permeabilized using detergents or alcohol, allowing antibody penetration into the cytoplasm (guide ATcfc, 2023). This approach is widely employed for the detection of cytokines, enzymes, transcription factors, and signaling pathway components. Of note, the use of viability dyes is critical to monitor live/dead discrimination during staining protocols. Dead cells can interfere with antibody binding and exhibit autofluorescence, ultimately leading to nonspecific labeling and spurious results (Bio Rad Antibodies, n.d.).

Flow cytometry is a technique widely employed in immunology and cell biology studies, enabling a detailed investigation of the immune response and specific cell populations with high accuracy (McKinnon, 2018). In parasitology, flow cytometry has gained increasing relevance both in research and in certain clinical applications. It can be applied to assess parasite load, monitor drug response, and even detect parasite resistance, with potential epidemiological applications in endemic areas (Grimberg, 2011).

Within the context of flow cytometry, it is also possible to implement assays known as Cytometric Bead Array (CBA), which allow the simultaneous quantification of a variety of analytes and, when compared to ELISA, for instance, substantially reduce sample volumes and the time required to obtain results (Medeiros and Gomes, 2019). CBA is based on coupling (binding) a capture molecule (antibody, peptide, protein, among others) to beads (microspheres) internally dyed with different fluorescence intensities, with the purpose of capturing the desired analyte in different samples. Detection and quantification of the analyte(s) are performed through the addition of specific detection antibodies conjugated to a fluorescent molecule for each analyte, which, after acquisition on the flow cytometer, will emit a fluorescent signal corresponding to the concentration of the analyte present in the sample (Morgan et al., 2004). Several studies have already described the use of CBA, such (Ker et al. 2019), which performed a multiplex assay using beads previously coated with two recombinant Leishmania antigens: rLci1A and rLci2B. When evaluated against canine serum samples with VL, the assay demonstrated high sensitivity and specificity in distinguishing diseased, vaccinated and healthy dogs.

In cases of cutaneous leishmaniasis, flow cytometry assays have been used in serological studies for diagnosis and treatment monitoring, showing lower specificity than ELISA but higher sensitivity (Pedral-Sampaio et al., 2016). For individuals with malaria, accurate and precise diagnosis is crucial to prevent the progression of the clinical condition of the disease. In this context, flow cytometry has been employed for the analysis of blood infection through the use of antibodies conjugated to fluorochromes specific for nucleic acid staining. Since parasites multiply within erythrocytes, and these cells do not contain DNA, fluorescence is only detected when it binds to the parasite's DNA inside the erythrocyte, thus distinguishing infected cells from healthy ones based on fluorescence intensity. Fluorescence intensity also increases as parasites multiply through mitotic divisions, and this can be analyzed to differentiate the developmental stage of the parasite in blood samples (Janse and Van Vianen, 1994).

In T. cruzi infections, flow cytometry has been applied in studies to support the serological diagnosis of Chagas disease, as well as in the follow-up of individuals after treatment with specific drugs, through the use of fixed and labeled T. cruzi epimastigote forms for the assessment of IgG reactivity in serum samples, demonstrating high sensitivity and specificity for diagnosis (Matos et al., 2011). Moreover, flow cytometry has also been used as an alternative serological approach for VL identification and as a tool to characterize the humoral response against Leishmania infantum in serum samples from VL-HIV coinfected patients (Matos et al., 2011) and presents a higher sensitivity and lower specificity in relation to indirect immunofluorescence in CL infection (Andrade et al., 2010; Rocha et al., 2002).

Among its advantages are high sensitivity and speed. However, it is a more complex and costly technique, requiring specialized equipment and trained personnel. Nonetheless, with the progressive reduction in the cost of cytometers, flow cytometry is expected to play an increasingly important role in the evaluation of parasitic infections, particularly in the development and validation of new drugs and immunotherapies (McKinnon, 2018; Grimberg, 2011). In summary, flow cytometry has increasingly emerged as a versatile tool with broad applicability in the diagnosis of parasitic diseases, enabling the detection and characterization of infections and contributing to significant advances in both research and clinical practice.

PCR is one of the most widely employed methodologies in diagnostic laboratories, as it enables the rapid generation of results that guide appropriate therapeutic decisions in response to detected infections. The process mimics the natural duplication of DNA in vivo, albeit with key differences, including the exclusive use of DNA polymerase (Khehra et al., 2025).

The reaction requires essential components: primers, DNA polymerase, PCR buffer, cofactors, deoxynucleotide triphosphates (dNTPs), and water. The thermostable enzyme responsible for synthesizing the DNA strand is Taq polymerase, originally isolated from the thermophilic bacterium Thermus aquaticus. Primers act as specific initiators for the selected target sequence. The PCR buffer provides an optimized environment, containing cofactors such as MgCl₂, while the dNTPs—thymine (T), adenine (A), cytosine (C), and guanine (G)—serve as the building blocks for DNA synthesis. When present in the correct proportions, these components ensure efficient enzyme activity and overall reaction performance (Garibyan and Avashia, 2013).

The PCR cycle consists of three major steps: denaturation at 94 °C −95 °C to separate the DNA strands; annealing at a reduced temperature to allow proper primer binding; and extension at 68 °C −72 °C, depending on the kit used, to synthesize the new DNA strand. An additional 5-min extension step at the end of the cycles is commonly included to ensure completion of amplification (Figure 7) (Khehra et al., 2025).

Conventional PCR requires that the patient's sample DNA be extracted using column-based methods, phenol/chloroform, or magnetic beads. It is worth noting that at low concentrations, there is an increased likelihood of false negatives and/or nonspecific results, in addition to the potential contribution of inhibitors to reaction failure, as well as interference from proteins and salts (Khehra et al., 2025). Therefore, this reaction relies on several factors to be properly executed. Its application in diagnostics is still employed, particularly in regions lacking the infrastructure to routinely perform different PCR modalities, since the resource requirements and higher costs of reagents for other techniques often make the implementation of newer methods unfeasible in more remote settings.

New PCR modalities have been developed, including conventional PCR, qPCR (Quantitative), Nested PCR, among others. Conventional PCR remains the most widely used in many laboratories due to its low cost and reliable amplification at a relatively affordable level, in addition to its ability to amplify larger targets. The method is followed by a qualitative analysis, as it indicates the presence or absence of a specific pathogen through the appearance of bands in electrophoresis corresponding to the gene within the target DNA (Garibyan and Avashia, 2013). Comparing to this, quantitative PCR, or qPCR, enables not only detection but also the estimation of absolute or relative quantification of the number of amplified DNA copies of the target gene (Arya et al., 2005).

Western blotting (WB), also known as immunoblotting, is a well-established and widely used technique in molecular biology and proteomics (Mahmood and Yang, 2012). This method allows the detection, identification, and quantification of specific proteins within a biological sample. The term “blotting” refers to the transfer of proteins, previously separated by gel electrophoresis, onto a membrane (typically nitrocellulose), where they are subsequently recognized by specific antibodies (Towbin et al., 1979; Burnette, 1981). WB is notable for its high sensitivity and specificity, enabling the detection of proteins even at very low concentrations (Kurien and Scofield, 2006). Beyond its widespread use in scientific research, Western blotting is also applied in the diagnosis of various diseases, including infections and genetic disorders (Mahmood and Yang, 2012).

According to (Heidari et al. 2019), Western blotting was evaluated for its diagnostic performance by detecting a specific pattern of immunodominant Leishmania infantum proteins using sera from patients with visceral leishmaniasis. The results showed that the technique could identify multiple proteins specifically recognized by patient antibodies, confirming its high sensitivity and specificity. Thus, Western blotting proved extremely useful for confirming suspected cases or those with inconclusive serological results, making it a valuable complementary diagnostic tool in laboratories with adequate infrastructure. As reported by (Ascanio et al. 2024), although Western blotting demonstrates high accuracy in confirming Trypanosoma cruzi infection, operational limitations—such as high cost, technical complexity, and limited availability—restrict its use to reference laboratories and research centers. In this context, the method is essential for resolving cases with discordant serological results but is not routinely implemented in conventional clinical laboratories.

For malaria diagnosis, this technique proved effective in detecting different Plasmodium species and has been considered a useful tool for serological diagnosis, mass screening in endemic regions, and safety testing in blood transfusions in areas with prevalent P. vivax malaria (Son et al., 2001). In the context of Chagas disease diagnosis, Western blotting has been described as a supplemental method, particularly useful when analyzing samples from children under 5 years old from various regions of Brazil (Frade et al., 2011).

In patients co-infected with other pathogens, especially Leishmania species, cross-reactivity often limits the specificity of serological tests, representing a major challenge in areas endemic for both T. cruzi and Leishmania spp. Although the use of recombinant antigens can partially reduce cross-reactions, it does not fully resolve the issue (Riera et al., 2012). Western blotting offers an advantage over conventional serological techniques because it allows the identification of antibodies targeting distinct polypeptide fractions within complex parasite antigen mixtures. Consequently, this approach can provide higher specificity and improved performance compared to other immunological methods (Riera et al., 2012).