Where Have All the Diagnostic Morphological Parasitologists Gone?

ABSTRACT

Advances in laboratory techniques have revolutionized parasitology diagnostics over the past several decades. Widespread implementation of rapid antigen detection tests has greatly expanded access to tests for global parasitic threats such as malaria, while next-generation amplification and sequencing methods allow for sensitive and specific detection of human and animal parasites in complex specimen matrices. Recently, the introduction of multiplex panels for human gastrointestinal infections has enhanced the identification of common intestinal protozoa in feces along with bacterial and viral pathogens. Despite the benefits provided by novel diagnostics, increased reliance on nonmicroscopy-based methods has contributed to the progressive, widespread loss of morphology expertise for parasite identification. Loss of microscopy and morphology skills has the potential to negatively impact patient care, public health, and epidemiology. Molecular- and antigen-based diagnostics are not available for all parasites and may not be suitable for all specimen types and clinical settings. Furthermore, inadequate morphology experience may lead to missed and inaccurate diagnoses and erroneous descriptions of new human parasitic diseases. This commentary highlights the need to maintain expert microscopy and morphological parasitology diagnostic skills within the medical and scientific community. We proposed that light microscopy remains an important part of training and practice in the diagnosis of parasitic diseases and that efforts should be made to train the next generation of morphological parasitologists before the requisite knowledge, skills, and capacity for this complex and important mode of diagnosis are lost. In summary, the widespread, progressive loss of morphology expertise for parasite identification negatively impacts patient care, public health, and epidemiology.

TEXT

Recent advances in molecular and proteomic-based laboratory diagnostics have revolutionized the detection of medically important parasites in humans and animals. These advances have enhanced global access to reliable diagnostics, facilitated the detection of common parasites, and enabled the identification and characterization of novel parasitic pathogens. For example, rapid antigen detection tests (RDTs) are now widely used for routine diagnosis of malaria throughout regions of endemicity and form an important component of the World Health Organization Guidelines for Malaria (1). Next-generation, highly sensitive malaria RDTs rival the sensitivity of microscopy. Additionally, nucleic acid amplification tests (NAATs) such as real-time PCR are now widely used for the detection of certain parasitic pathogens such as Toxoplasma gondii and Trichomonas vaginalis and are prominently featured in national diagnostic guidelines (2, 3). Notably, the expanded adoption of multiplex NAATs for human gastrointestinal infections has enhanced the identification of common intestinal protozoa in feces, while allowing for the simultaneous detection of common bacterial and viral pathogens (4). Most recently, unbiased “shotgun” metagenomic next-generation sequencing (mNGS) has emerged as a promising diagnostic tool for detecting all classes of potential pathogens in various clinical specimen types. This method has allowed for the detection of unsuspected cases of toxoplasmosis (5) and granulomatous amebic encephalitis (6) in settings where conventional diagnostics were unrevealing. Additional methodologies such as proteomics (e.g., mass spectrometry) and flow cytometry also show promise for the identification of select parasites (7, 8).

Despite the benefits of advanced parasite diagnostics, traditional microscopy-based morphologic analysis remains the gold standard diagnostic method for many parasitic infections and continues to be the most appropriate, cost-effective, and sometimes, the only accurate way to identify parasites in many settings. Thus, it is concerning that increased reliance on nonmorphology-based detection methods has contributed to the progressive, widespread loss of microscopy skills and morphology expertise for parasite identification. Loss of microscopy and morphology skills has the potential to negatively impact patient care, public health, and epidemiology. Missed and inaccurate diagnoses can lead to poor patient outcomes, inappropriate treatment approaches, and mischaracterization of potential pathogens. This gap in skills and expertise is not easily bridged by modern diagnostic techniques. Although attempts have been made to train artificial intelligence (AI) to analyze smears, expert validation of AI-derived results is still required before final reporting (9–12).

Unfortunately, microscopy and morphology skills are not easily replaced because it takes several years of training to become an effective parasite morphologist. Parasitology is inherently challenging as it represents applied zoology and may require different approaches and thought processes from those commonly taught and reinforced in medical laboratory training programs. Specifically, morphologic diagnosis requires significant practical and theoretical knowledge of anatomy, biology, zoology, taxonomy, and epidemiology across the vast array of parasite taxa capable of infecting humans. Morphological expertise cannot be replicated without active and sustained attention to training a new generation of morphologists.

In this commentary, we emphasized the importance of maintaining competencies in microscopy and morphological parasitology skills within the medical and scientific community. The following sections address the limitations of nonmorphology-based methods and the consequences of losing morphologic skills on patient and public health. We concluded with recommendations for training the next generation of diagnostic parasitologists before the requisite knowledge, skills, and capacity for this complex and important mode of diagnosis are lost.

LIMITATIONS OF NONMORPHOLOGY-BASED TESTING

Non-morphology-based diagnostics are not available, appropriate, or suitable for the detection of all human and animal parasites in all settings. Table 1 addresses the strengths and limitations of nonmorphology-based parasite diagnostics, and these are discussed in greater detail below.

TABLE 1.

Strengths and limitations of morphology- and nonmorphology-based parasite diagnostics, (not all-inclusive)^a

Diagnostic test characteristic	Morphology based diagnostics	Serology based diagnostics	PCR based diagnostics	Sequencing based diagnostics
Sensitivity	++	+++	+++	+++
Specificity	+++	+	+++	+++
Quantification	+++	+	++	−
Turnaround time	+++b	++	++	+
Cost-effectiveness	+++	++	++	+
Technical complexity	+	++c	+++	+++
Objectivity of results	+	++	+++	+++
Genus level identification	+++	++	+++	+++
Species-level identification	++	+	+++	+++
Genotype level identification	−	−	−	+++
All parasites detected in one test	+++	−	−	−d
Capacity to detect novel or zoonotic not previously described agents	+++	−	−	+++
Variety of specimen matrices suitable for testing	+++	−	+++	+++
Easily adapted for use in the field and resource-poor settings	+++	−e	−	−
Capacity to detect acquired drug resistance	+	−	++	+++

^a−, no capacity/efficacy; +, limited capacity/efficacy; ++, moderate capacity/efficacy; +++, high capacity/efficacy.

^bWith the exception of histology slides and permanent stained fecal smears.

^cWith the exception of rapid antigen/antibody tests which are low complexity.

^dA recent high-throughput sequencing-based protocol has successfully achieved this for blood parasites (16, 17).

^eWith the exception of rapid antigen/antibody tests, which are easily adapted to low-resource and field settings.

Insufficient coverage of medically important parasites.

First and foremost, it must be noted that commercial and laboratory-developed tests do not exist for all medically important parasites. Most available are designed specifically to detect a limited number of known species. Humans are known to harbor at least 848 species of helminths, protozoans, and arthropods; of these, about 90 species commonly infect them (13–15). NAATs are only available for a few of these parasites even in most of the advanced clinical laboratories, viz., Plasmodium species, Entamoeba histolytica, Giardia duodenalis, Cryptosporidium spp., Cyclospora cayetanensis, Toxoplasma gondii, Trichomonas vaginalis, some soil-transmitted helminths. The DNA-based identification of arthropods is mostly limited to research and public health surveillance programs, while clinical diagnostic laboratories routinely receive whole samples of insects and arthropods for identification. Most routinely employed multiplex PCR assays are limited to common protozoal parasites and may not detect nor identify the less common, genetically dissimilar, or emerging parasitic agents. This is particularly problematic in laboratories that no longer diagnose clinical fecal samples by microscopy because infection with helminths and protozoa not in the array target panels of such commercial multiplex PCR assays will not be diagnosed and were, therefore, not considered for treatment indications. Furthermore, commercial multiplex PCR assays are not yet validated for use in environmental samples (food, water, and sewage samples). Attempts to fill these gaps by commercial diagnostic companies have been hampered by the unavailability of morphologically well-characterized specimens for test validation, thus emphasizing the importance of maintaining microscopy skills.

Unfortunately, attempts to develop molecular-based assays that will identify all potential parasitic pathogens in a sample have met with limited success. A targeted amplicon deep sequencing approach has been developed to detect all blood parasites in a single blood sample, but the small size of the amplified target does not allow for species-level differentiation of all human infecting filariae (16, 17), a task which can be achieved by microscopy. Similarly, attempts to use metagenomics or shotgun mNGS to detect and identify all parasites in fecal samples have had limited success (6, 18). Such approaches are high-cost and often provide little clinical gain compared to the microscopic identification of parasitic agents. In comparison, expert microscopic examination applies to all parasites.

Specimen incompatibility for molecular testing.

Even when suitable molecular testing is available, the extraction of high-quality nucleic acid and subsequent molecular analysis may not be possible in all situations. Feces particularly presents a challenge for PCR diagnostics due to the presence of inhibitors, such as bile salts, urates, complex polysaccharides, stercobilinogen, and stercobilin (19). While advances in fecal DNA extraction methods have reduced the prevalence of inhibition, it remains a relevant consideration in the diagnostic screening of fecal specimens for parasitic infection.

The greatest challenge in this context is the widespread formalin fixation of specimens. This also affects the use of NAAT methods in samples prepared for histopathological diagnosis, where the routine use of formalin as a fixative before tissue embedding rapidly degrades DNA (20), making it difficult to extract unfragmented DNA for subsequent molecular analysis. Arthropods and adult helminths are also often preserved in formalin after specimen collection or upon receipt in frontline laboratories, and thus may be less amenable to molecular testing. To avoid issues with further molecular testing, samples should be divided into aliquots where possible, one aliquot stored without any additive or placed in sterile physiological saline, and one placed in a suitable fixative. There are now several commercially available fixatives that preserve fecal parasite morphology while still allowing DNA extraction. If this is not possible, storage in 70% pure ethanol is preferable (note that denatured ethanol may contain PCR inhibitors).

Inadequate sequence reference databases.

In cases where DNA extraction and sequencing of a parasitic pathogen is possible, sequence analysis and accurate identification still require the availability of inclusive reference databases. Regrettably, sequence data for all parasite species that infect humans are not currently available, and this may result in a missed or incorrect diagnosis of potentially novel or important agents. For example, a case of microscopy confirmed that Mansonella perstans could not be definitively detected in blood using shotgun metagenomic NGS, despite employing four different bioinformatics tools/databases (21). Another example is the initial announcement that a Guinea worm (Dracunculus medinensis) had been detected in a patient in Vietnam, who had no travel to areas of endemicity, based on the closest match in GenBank. Upon morphologic characterization, the Dracunculus specimen was found to be inconsistent with D. medinensis. However, further sequencing could not definitively resolve the potentially novel species due to a lack of reference sequences. While this situation will undoubtedly improve over time as more high-quality sequence data are added to reference databases, the current situation poses significant limitations on the broad use of sequencing for parasite detection and identification. Additionally, reference guidelines for standardized and internationally comparable molecular sequencing of parasitic agents, such as those available for bacterial sequencing through the Clinical and Laboratory Standards Institute (22, 23), were not available.

Cost and complexity of molecular testing.

In general, molecular tests are significantly more expensive and complex than traditional microscopy, due to the need for specialized instruments, reagents, and workflows. This greatly limits their widespread implementation, particularly in resource-poor and field-based settings where stable electricity, refrigeration, and dependable supply chains are not widely available. Even in resource-rich settings, the high cost of some molecular tests such as targeted and shotgun metagenomic NGS testing may be cost-prohibitive. If used for routine diagnosis, they could place a significant burden on the health care system.

Inability to differentiate active from past infection.

A limitation of serological diagnostic methods is their general inability to differentiate active from past infections due to the persistence of IgG antibodies following treatment (24). For antigen-based tests, persistent antigenemia in HRP2-based malaria RDT positivity can occur in up to 50% of cases 1 month posttreatment and has been documented up to 2 months posttreatment (25). A similar problem with prolonged false positives may be observed in PCR testing of some tissue-dwelling parasites species, particularly schistosomes, due to the persistence of circulating parasite antigens and nucleic acids from nonviable parasites for up to 135 days following treatment (26).

In some cases, microscopy allows for the assessment of parasite viability and motility, which may support the assessment of active infection. This assessment holds value for the evaluation of certain parasitic diseases such as schistosomiasis. For example, microscopy can distinguish between viable and nonviable Schistosoma eggs (27), which has implications for correct patient management, disease course, and public health control.

Turnaround time.

Unlike many microscopy analyses, molecular testing is not typically performed around the clock, and specimens are frequently tested in batches to save on the cost of controls. Therefore, some settings may see significant delays in molecular testing results that are not seen with routine microscopy. This is particularly true if testing is not performed in-house and must be sent to a reference laboratory. While factors such as choice of test and specimen processing bottlenecks may delay microscopic testing, the time taken for simple microscopy by an experienced operator, even with prior concentration techniques, remains swifter than DNA extraction and analysis by PCR.

Need for multiple analyte-specific tests.

A limitation of targeted molecular testing, as well as serology and antigen-based detection methods, was the need for clinical suspicion to guide test ordering. Even with a narrowed differential diagnosis, multiple PCRs or serological tests may ultimately be required to make the diagnosis, and even these may not detect uncommon parasites. In comparison, microscopic examination can identify all morphologically intact parasites in a specimen.

Unreliable results.

Test reliability may vary due to multiple factors, including the quality of the test, storage conditions of the reagents, and the state of the equipment. While this may also apply to microscopy (as discussed below), there are some specific examples of nonmorphologic parasite tests that deserve mention. For example, malaria RDTs, are widely used but may frequently yield both false positive and false negative results (28). Possible reasons include misinterpretation of the test line pattern in any setting, poor storage conditions, or shelf life of the RDTs in resource-limited settings. For this reason, malaria control guidelines recommend RDTs only when quality-assured microscopy is not available, and confirmation by microscopy for RDT-positive samples before starting the anti-malarial treatment (1). It is also important to note that malaria RDTs will only provide a qualitative result for the targeted Plasmodium species. Expert malaria microscopy is still necessary for determining the stage and species of the parasite in the blood, calculating the parasite density, and detecting concurrent, nonmalarial diseases such as relapsing fever borreliosis (1). Malaria RDTs are also not equipped to detect the various mutations and polymorphisms in the targeted genetic regions or the expressed protein. The emergence and rapid spread of HRP2/3 diagnostic antigen gene mutations in Plasmodium falciparum have quickly rendered many rapid antigen tests for falciparum malaria, which specifically target these antigens, less reliable (29). Given the potential mortality of severe malaria, case reports of travelers returning with pfhrp2/3 deletion P. falciparum malaria requiring rapid microscopic diagnosis in the context of failed RDT tests (30) raise concerns. Using a combination of HRP2/3-based and LDH-based RDTs may resolve this issue, but the increased cost will usually prevent the implementation of this strategy in low-resource settings.

Molecular tests are also not without limitations. In some cases, PCR assays are generic and will not differentiate rare parasitic species from common pathogens, for example, the most used Strongyloides stercoralis real-time PCR is genus-specific and does not differentiate infections with other Strongyloides species (31, 32). Some PCRs may be used to specifically detect E. histolytica sensu stricto and differentiate it from morphologically identical non-pathogenic members of E. histolytica complex. However, some commercial PCR assays, such as the BioFire FilmArray Gastrointestinal panel, will result positive for the potentially pathogenic E. histolytica when the non-pathogenic E. dispar is present in large concentrations and, therefore, provide little advantage over microscopy in terms of specificity.

Similarly, many serologic tests are not designed for or capable of species-level distinction, despite the unknown effects this may have on sensitivity and specificity (33–35). Diagnostic serology for parasites is a challenging area of research, and many routinely used serological assays may yield false-positive results due to the formation of cross-reactive antibodies to other parasites. In some cases, such as the diagnosis of S. stercoralis infection (32) and E. histolytica liver abscess in those without prior residence in higher regions of endemicity (36), serology has high utility. In other uses, such as the serological diagnosis of Toxocara and Taenia solium (cysticercosis) infections using specific commercial assays, cross-reaction with other parasites may confound results (37, 38). The development of new and specific recombinant antigens, and their use in duplex assays may address some of these challenges.

Insufficient utility for describing new taxa.

Despite advances in molecular testing, morphological characterization is still required for the valid description of the key anatomical features of a new species of parasite, even if DNA sequences have already been derived (39). The naming of a new species based entirely on sequence data without a sufficient morphological description of the type specimen(s) results in a nomen nudum. Such names are considered invalid by the code set forth by the International Commission on Zoological Nomenclature (40). For example, reports of human infections with “Candidatus Dirofilaria hongkongensis,” also sometimes described as “Dirofilaria hongkongensis” are nomen nudum as the original paper using this species epithet did not include either a morphological description or provide a holotype (41). The term “Candidatus” was approved by the International Code of Nomenclature of Prokaryotes (ICNP) for the description of new prokaryote taxa for which molecular evidence, but incomplete further data for full taxonomic description, were available. It has not been approved or accepted by the ICZN (41). This term is now increasingly, though incorrectly, creeping into the parasitology literature for the description of new parasite taxa that have been defined by molecular means only, consequently compromising the literature and our definitions of infecting species. While molecular methods have a growing role in the diagnosis of parasitic infections, they should complement rather than replace the morphological assessment of 'new' parasites (e.g., zoonotic) or where morphology is of limited utility (e.g., degraded or distorted organisms in histology sections).

IMPACT OF INADEQUATE MORPHOLOGY SKILLS

The sections above discuss the limitations of nonmorphologic diagnostics and highlight the need for the maintenance of expert parasite morphology skills. We propose that there is a recognized gap in available morphologic expertise in laboratory medicine, especially in industrialized settings where parasites are uncommonly encountered. Certain modified morphology-based methods (e.g., autofluorescence of coccidian protozoans) or education practices (i.e., using clinical manifestations and travel history to assist with morphological identification) may partially overcome the issues associated with inadequate morphology skills; however, the definitive diagnosis is often suspected or made using basic morphology. The consequences of inadequate morphology skills are many, including missed or inaccurate identifications, which can have significant implications for individual patients, and may confound surveillance data, impacting the delivery and limited resources of disease control programs.

Parasite misidentifications.

Examples of misidentifications in the medical literature are numerous and include reported cases of parasites occurring where they are known not to occur without supporting morphological or molecular data to support the identifications given. For example, there are multiple reports of loiasis (Loa loa) from India, a non-region of endemicity, where the vector is not known to be present. These most probably represent misidentifications of zoonotic Filarioidea (42–46). Similarly, there are recent reports of dracunculiasis (Dracunculus medinensis infections) which may be due to Dirofilaria spp., Spirometra spp., other parasites, nonparasitic pathological processes, or other artifacts, but which cannot definitively be identified as D. medinensis by the information reported (47–51). Additionally, there have been reports of Gnathostoma spinigerum from Latin America (52), where only Gnathostoma binucleatum is currently known to infect humans (53) and where G. spinigerum is not present (54). There have also been recent reports of Paragonimus westermani from West Africa (55, 56) where this species does not occur but P. africanus and P. uterobilateralis are endemic (57).

In other cases, the species epithet assigned to a parasite is the most common or most well-recognized species infecting humans, rather than the actual species infecting the patient. This often occurs because generic serological or molecular tests are incorrectly reported as species-specific, or due to the use of outdated references. Such reports introduce errors into the literature which may be subsequently cited and retained, or incorrect sequences are deposited in online databases. This matters because in many cases different species of the same genus have different intermediate and reservoir hosts, clinical presentation, and geographical ranges. Cases of parasites being incorrectly assigned to the commonly known species such as P. westermani, G. spinigerum, Trichinella spiralis, and Dibothriocephalus latus (syn. Diphyllobothrium latum), despite being other species of the same genus or family, are not uncommon. In some geographic regions, “rare” parasitic diseases may be more common than generally appreciated. For example, cases of babesiosis in Africa and China have been misdiagnosed as malaria (58, 59). Similarly, infections with Oesophagostomum bifurcum, Ternidens deminutus, Strongyloides fuelleborni fuelleborni, Strongyloides fuelleborni kellyi, and Trichostrongylus spp are characterized by the passage of eggs commonly misidentified as hookworm infections by inexperienced operators (60–64). The prevalence of opisthorchiasis and clonorchiasis in some regions is also likely vastly inflated by misidentification due to superficial similarities with other fish-borne trematodes (especially other heterophyid trematodes) (65). When it comes to arthropods, the lack of awareness and loss of morphological identification skills often results in genus-level identifications of important vectors and pathogens or generic identifications such as “maggot” or “fly.” An additional challenge with arthropods is even determining which may be clinically relevant and which represent an incidental finding or environmental contamination of a clinical specimen.

An example of the effect that the loss of morphology skills may have on parasitic disease control programs, strongyloidiasis has been recently included by the WHO among the targets of the Neglected Tropical Diseases NTD Roadmap to 2030 (66). Accurate differentiation between S. stercoralis, hookworm, and other hookworm-like helminth eggs (32, 61, 63, 67) will be required for accurate mapping, implementation, monitoring, and evaluation of the control program should classic parasitology methods based on the retrieval of parasite larvae be recommended for the diagnosis of strongyloidiasis, such as Baermann technique (68). Additionally, the validation of effective serological tools for S. stercoralis surveillance applications will rely on the accurate characterization of cases via agar plate culture or Baermann sedimentation as a reference standard.

Such mistakes confound the scientific literature and introduce ongoing errors regarding the geographical distribution, clinical presentation, biology, and epidemiology of parasitic diseases. Additionally, genetic sequences associated with inaccurately identified parasites may be uploaded to DNA sequence databases, further perpetuating incorrect identifications in subsequent publications (69–72). This problem has previously been recognized in other fields of taxonomy as well (73). In some cases, parasite sequences have been incorrectly included in published whole-genomes of their host species, a potential confounder in sequencing-based identification (74).

Misinterpretation of nonparasitic findings.

Inadequate morphology experience has also led to inaccurate reports of nonparasitic pathological processes and artifacts as parasites, and in some cases, has resulted in erroneous descriptions of new human parasitic diseases. In some instances, entirely new parasites of humans have been described and this error has then been reproduced multiple times in the scientific literature (75–80). For example, a brief literature search by the authors identified over 30 manuscripts describing case reports or case series of Lophomonas blattarum (a parasite of cockroaches) infecting the human respiratory tract. All these reports likely represent misidentifications of bronchial ciliated epithelial cells, a motile constituent of the human respiratory tract referred to as ciliocytophthoria (81).

Loss of morphology skills led to a compromised review process.

There is a great increase in the submission and publication of parasitic disease cases to journals at a time when there has been a great decrease in the availability of those with the experience to expertly review such cases. It is becoming increasingly difficult for such errors to be identified before publication by journals. Few experienced parasite morphologists are available to review these manuscripts, while many potential reviewers who are currently active in parasitology research do not have significant morphology experience. Erroneous identifications of parasites are particularly an issue in medical discipline-specific journals (e.g., dermatology, ophthalmology), where editorial boards and the reviewer pool generally lack parasitology training beyond the standard (limited) medical curriculum. In such cases, it is crucial to invite reviewers with expertise in clinical parasitology to ensure accurate reporting. Unfortunately, by the time a publication is prepared, the patient will have long since been treated (potentially incorrectly) or discharged. Therefore, published case reports must describe all diagnostic practices in enough detail that any errors in diagnosis may be identified by reviewers and readers.

An additional issue is that authors of case reports often exclude the laboratory parasitologist who made the initial diagnosis in the authorship process, meaning that insufficient data on how the identification was arrived at is available for scrutiny. We feel that case reports should not be published unless all relevant morphological and morphometric data that was used to arrive at the identification given is provided. This is compounded by the proliferation of medical journals over the past 2 decades, the emergence of many “predatory journals” with low standards of scientific and editorial review, and increased referencing of unreviewed preprints. Furthermore, the increased workloads placed on experienced morphological parasitologists in diagnostic and public health laboratories have reduced their available time for review and editorial activities.

CONCLUSIONS AND A WAY FORWARD

Expertise in parasite morphology diagnosis will remain essential in all healthcare settings for decades to come, both in high- and low-resource settings. While the prevalence of some parasitic diseases has decreased in the last few decades, other parasitic diseases previously considered rare have increased in prevalence. Primate malaria is currently emerging as a human infection in several parts of the world (82–85). Other rare and zoonotic parasitoses that may be unfamiliar to laboratorians include dirofilariasis (86), zoonotic onchocerciasis (87), gnathostomiasis (88, 89), alveolar echinococcosis (90, 91), Ancylostoma ceylanicum (92) and Ancylostoma caninum hookworm disease (93), angiostrongyliasis (94), and acanthacephaliasis (95). While PCRs for these pathogens have now been developed, they highlight the need to screen for pathogens that are not always considered in routine diagnostic PCR panels.

Without concerted action by the medical and scientific community, the issue of limited availability of parasitic morphology diagnostic expertise will become more pressing in the future. Many current morphologists working in reference laboratories are reaching retirement age and few younger parasitologists are being trained in their place. With time, there is also significant attrition in the laboratory techniques and infrastructure required for the morphological diagnosis of parasites. Detailed, comprehensive morphological keys are often only present in old publications and are not easily retrievable or accessible unless “inherited” from mentors. When those diagnosticians do train new clinical microbiologists, they generally find employment in other laboratories, where their parasitology skills and proficiency cannot be maintained. In industrialized nations, morphological parasitology largely has been minimized or excluded from undergraduate medical and medical laboratory technology, pathology residency, infectious disease, and clinical microbiology fellowship training curricula. Globally, morphology expertise, despite its unique value and the gold standard for parasite diagnosis, is often not as highly regarded, nor attended to, as molecular skills among early career scientists, medical technologists, and clinical microbiologists, despite requiring far greater depth and breadth of knowledge. The inclusion of “innovation scores” in large-scale government grants biases researchers toward more modern and expensive molecular techniques rather than the use of traditional morphology, even in cases where the latter may be more or equally suitable to fulfill the purposes of the intended research. The highest burden of parasitic diseases is in low-income countries, meaning that government funding for the development and maintenance of parasitology diagnostic and research skills is a low priority relative to other conditions in those nations most able to afford such funds. This lack of available funding opportunities is leading to a loss of morphology expertise, hence lowering its contribution to the diagnostic process, with negative implications for clinical and veterinary parasite diagnosis.

To address these challenges, we recommend active measures be taken to increase the teaching of morphological parasitology in graduate and postgraduate degrees and to improve the value and importance placed on these skills in the wider clinical microbiology community (Table 2). Both in low- and high-resource settings, the value of, and need for, well-trained parasite morphologists and their value in the field of clinical parasitology should be emphasized, to motivate the next generation. The introduction of newer techniques was not meant to supplant morphological techniques, but rather should coexist with them. Hence there is a need for correct sampling and collection guidelines, allowing for both techniques to be performed. It is important to retain the discipline of morphological parasitology in the laboratory and to consult those who practice it, particularly in the context of the diagnosis and description of new or rare species infecting humans. Its loss will lead to errors in science and poorer outcome for those who matter the most, our patients.

TABLE 2.

Tips for improving parasitological skills

Tip	Examples and Principles
Strive to go above and beyond	When encountering an interesting case, go out of your way to learn everything you can about that organism.
Don’t be afraid to make a mistake	None of us are infallible. If you misidentify something, take it as a learning experience. Learning more about the organism will limit misidentifications in the future.
Archive positive specimens and keep an image library	Keep positive specimens and take images of positive cases you encounter. These are great for teaching and maintaining competency and proficiency. They can also be helpful for networking and possible publication opportunities (see below)
Seek out additional education in the discipline	•If continuing education units (CEUs) are required in your job, seek out those specific to parasitology.•Review the CDC’s DPDx monthly case studies: (https://www.cdc.gov/dpdx/monthlycasestudies/2020/index.html).•Review the accompanying reports and educational material for parasitology proficiency testing/external quality assessment schemes (e.g., through the CAP, API, UKNEQASa).•Attend hands-on workshops, take classes or obtain a degree in medical parasitology (e.g., through a reputable school of tropical medicine).
Attend society educational meetings	This can be challenging due to budget restrictions but attending meetings can be a great way for learning what’s new in the field and for networking. Two of the best for human clinical parasitology in the United States are the annual meetings by ASM (Microbe) and the ASTMH. In Europe, there is ECCMID organized by ESCMID. ESCMID has a Study Group for Clinical Parasitology (ESGCP). The Australian Society for Microbiology has a Parasitology Special Interest Group, and the ASP holds annual scientific meetingsa.
Keep up on the literature	This can also be challenging, especially because publications in parasitology transcend the medical, veterinary, and zoological communities, but it is a great way to keep up with new diagnostic methods and nomenclatural changes. Image-based challenges are particularly used in testing one's morphology skills as they often show classic morphology-based preparations (e.g., JCM, CID, CMI, and NEJM Image Challengesa).
Publish in the scientific literature	If an interesting case is encountered, reach out to your colleagues, supervisor, laboratory director, and experts in the field about the possibility of a joint publication.
Networking – make yourself visible	1.Join appropriate listservs (e.g., Division-C and ClinMicroNet hosted by ASM), and maintain an active presence; if someone presents a case, share your thoughts, and do not be afraid about getting it wrong (see above). 2. Be active on social media platforms, for example, Creepy Dreadful Wonderful Parasites (https://parasitewonders.blogspot.com/). 3. Share interesting cases with experts in the field – it shows us who is enthusiastic about parasitology 4. Join committees (e.g., ACTM, ASCP, ASM, ASM SIG, ASP, ASTMH, BSP, CAP, ESGCP, RSTMHa).
Ask for help	CDC DPDx, listserves, colleagues in the field.

^aACTM, Australasian College of Tropical Medicine; API, American Proficiency Institute; ASCP, American Society for Clinical Pathology; ASM, American Society for Microbiology; ASM SIG, Australian Society for Parasitology and Tropical Medicine and Parasitology Special Interest Group; ASP, American Society for Parasitology and/or Australian Society for Parasitology; ASTMH, American Society of Tropical Medicine and Hygiene; BSP, British Society for Parasitology; CAP, College of American Pathologists; CDC, U.S. Centers for Disease Control and Prevention; CID, Clinical Infectious Diseases; CMI, Clinical Microbiology, and Infection; ECCMID, European Congress of Clinical Microbiology and Infectious Diseases; ESCMID, European Society of Clinical Microbiology and Infectious Diseases; ESGCP, ESCMID Study Group for Clinical Parasitology; NEJM, New England Journal of Medicine; JCM, Journal of Clinical Microbiology; RSTMH, Royal Society for Tropical Medicine and Hygiene.

MAIN TAKE-HOME POINTS

•Basic light microscopy skills should be taught in all laboratory-based specialties, including the use of Kohler illumination and polarizing filters.

•Instructors should teach morphologic features, life cycles, and established endemic distributions of important human and veterinary parasites, in addition to information on clinical presentation and treatment.

•Journals should consult experts in parasitology when taxonomic changes or new names are being described.

•Good quality illustrations with defining features labeled must be provided to back up claims in refereed journal articles and conference presentations.

•Schools of tropical medicine and public health should maintain expert clinical laboratory referral and teaching units.

•Laboratories should participate in parasitology external quality assessment schemes.

•Ensure that the skills of expert parasitologists are valued (e.g. by including them in the authorship list of publications).

•Plan well in advance of their retirement for a succession of expert parasitologists, with ample time for training replacement staff.

•Laboratory leadership must ensure that the skills of expert parasitologists are valued, and plans made for succession, with ample time for training replacement staff, in advance of their retirement.

The views expressed in this article do not necessarily reflect the views of the journal or of ASM.