JoVE Journal > Immunology and Infection
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A Simple Fecal Flotation Method for Diagnosing Zoonotic Nematodes Under Field and Laboratory Conditions

Published: December 15, 2023
doi:
10.3791/66110
, , , ,
1Parasitology Department, Facultad de Medicina Veterinaria y Zootecnia,Universidad Nacional Autonoma de Mexico, 2Instituto de Biología,Universidad Nacional Autonoma de Mexico

Özet

This work describes the use of a flotation method to identify Toxocara canis and Ancylostoma spp. detected in fecal samples collected from dogs in Mexico from 2017 to 2021 under field conditions.

Abstract

Diagnosis of canine parasites with zoonotic potential such as Toxocara canis and Ancylostoma caninum under field conditions is usually difficult due to limited access to a laboratory in rural and suburban areas in Mexico. This study aimed to detect T. canis and Ancylostoma spp. in fecal samples collected from dogs in Mexico from 2017 to 2021 under field conditions. Sample size calculation resulted in a target enrollment of 534 dogs across the country.

Samples were collected directly from the rectum or the ground after defecation. Samples were stored in individual, tightly sealed, plastic bags at 4 °C. A saturated solution of sodium chloride (specific gravity [SpG] 1.20) was prepared both under field and laboratory conditions. Within 3 days of collection, 2-4 g of feces were tested for parasites using a flotation method by suspending each fecal sample in a saline solution. Feces were mixed with the flotation solution and crushed using a metal spoon.

Once a uniform consistency was achieved, the fecal sample was poured into a new plastic cup using a sieve and allowed to sit for 10-15 min. Three drops from the top of the mixture were collected using a sterilized inoculating loop. The slides were placed on the microscope and parasites were identified by trained parasitologists. Fecal samples from 1,055 dogs were screened microscopically. The number of positive samples for Ancylostoma spp. was 833 (78.95% frequency) and 222 (21.04%) for T. canis. These findings illustrate the importance of identifying zoonotic helminths in dogs living in urban and rural areas in Mexico using a coproparasitoscopic technique in the laboratory and under field conditions.

Introduction

Gastrointestinal parasites are one of the most common health problems that affect dogs1. Estimates suggest that there are ~700 million domestic dogs worldwide, and approximately 175 million may be categorized as free-roaming2. More than 60 parasite species are shared between dogs and humans, suggesting that dogs could be a source of infection for humans with these parasites3. Toxocara canis and Ancylostoma caninum are two parasitic species that infect dogs and, accidentally, human hosts. Currently, there are several studies about the locations where these helminths are able to survive and reproduce in Mexico. The prevalence of Toxocara in dogs varies from 0% to over 87% across the United States, Mexico, Central America, and the Caribbean4. Toxocara canis and Ancylostoma spp., as well as other parasitic species in dogs, have been previously reported in Mexico5,6,7,8,9,10,11,12,13 (Table 1).

Parasitic species Region Prevalence (%) Reference
Ancylostoma caninum Querétaro 42.90 5
Tabasco 15.90 6
Campeche 35.7 – 42.9  7
Yucatán 73.8 8
Babesia Morelos 13.60 9
Veracruz 10.00
Coccidial oocysts Yucatán 2.30 8
Ctenocephalides Morelos 30.3 10
Dipylidium caninum Yucatán 2.30 8
Dirofilaria  Yucatán 7.0 – 8.3 11
Giardia Tabasco 3.00 6
Yucatán 18.8 8
Leishmania Chiapas 19.00 12
Tapeworms Baja California 6.79 13
Toxocara canis Querétaro 22.10 5
Yucatán 6.20 8
Trichuris vulpis Yucatán 25.40 8
Trypanosoma Jalisco 8.10 9
Campeche 7.60
Chiapas 4.5 – 42.8
Quintana Roo 20.1 – 21.3
Toluca 17.50
Yucatán 9.8 – 34

Table 1: Regional prevalence (%) of dog parasites in Mexico from 2001 to 2020. Findings from previous investigations conducted from 2001 to 2020 have enabled the identification of canine parasite distribution across several urban and rural settings in Mexico. These studies provide a deep understanding of epidemiological elements conducive to the persistence of canine parasites in different ecosystems, contributing to a comprehensive assessment of the zoonotic impact of some parasite species.

Life cycle stages of intestinal parasites, such as eggs, cysts, oocysts, or larvae can be found in stool samples. Thus, examination of fecal material provides valuable information about the parasites of an animal. The need for a method to detect Ancylostomidae eggs in human feces led to the use of the simple fecal smear in 1878, which was used for many years to detect gastrointestinal parasites but was considered not very sensitive. Thus, the need arose to develop better copromicroscopic methods14. More than 100 years have passed since the flotation technique for recovering and counting parasite eggs in fecal samples was first described15. Since then, several methods and variants of the flotation technique have been considered a standard for the detection of some parasites in their hosts.

For example, Lane described a method in 1924 involving the direct centrifugal flotation technique, which integrates centrifugation followed by floating the sediment in a saturated sodium chloride solution with SpG 1.2 in 1 g (Lane) or 10 g (Stoll's modification). The flotation technique was subsequently modified by using solutions with different SpG14. In 1939, Gordon and Whitlock reported the disadvantages of Stoll's technique due to interference from detritus in visualizing parasite eggs and developed the quantitative method known as McMaster16. In 1979, O'Grady and Slocombe demonstrated that the specific gravity of the solution, timing, and mesh sizes of strainers affect the accuracy of egg detection using the flotation technique17. During the last decades, because several modifications have been made to the flotation technique, there is an urgent need for standardization of flotation methods. Currently, detection of canine helminth infections in the context of prevention of zoonotic parasites is required to apply appropriate anthelmintic treatments to limit environmental contamination with infectious stages of zoonotic nematodes18.

Among qualitative methods, the fecal flotation technique is widely used and accepted because it does not require much equipment, is simple, inexpensive, and reproducible; yet it has a major drawback in that it lacks sensitivity when the intensity of the infection is low19. The ability to reveal the presence of a greater number of parasitic elements such as eggs, oocysts, cysts, or nematode larvae is usually determined by the density of the solution20.

Previous reports have compared coproparasitological techniques for the detection of canine nematode eggs. With regard to the detection of motile protozoa, direct fecal smears are used; whereas sedimentation methods are useful for diagnosing heavy eggs of parasites such as trematodes21. One of the most widely used field-based diagnostic tests is the fecal smear method. However, the low level of sensitivity of this technique may be attributed to the fact that it does contain debris that interferes with the detection of parasite eggs. By incorporating a sieving step along with solutions that provide the proper SpG, the flotation method offers a clearer and less cluttered observation of ascarid and hookworm eggs. This leads to a more precise and efficient process for microscopic screening22. Likewise, simple flotation and direct centrifugal flotation techniques are very commonly utilized to recover parasite eggs and oocysts14. The classic flotation methods can be considered qualitative or quantitative depending on the use of a counting chamber such as the McMaster method15. Nonetheless, as the flotation technique has low sensitivity and focuses on the detection of parasites in the patent period, negative results should not be considered conclusive. However, the accuracy not only depends on the preservation procedure of fecal samples or the SpG of flotation solutions but also depends on the technical proficiency and experience in conducting fecal examinations of the user.

Consequently, other methods have been explored for the detection of canine parasites in feces. It has been generally recognized that one of the most widely used approaches for the diagnosis of intestinal helminth infections in dogs is the FLOTAC technique, a multivalent, sensitive, and accurate method that yields accurate and reliable results for the diagnosis of A. caninum in dogs when compared to a flotation protocol in a tube and the McMaster technique19,23. Sedimentation methods are useful for recovering fluke eggs, embryonated nematode eggs, and most tapeworm eggs, which cannot be recovered on the surface of a flotation solution because these structures do not float24. One method that has been proven to be superior to flotation/sedimentation techniques is the Modified Double Centrifugal Flotation method, as it enables the detection of cestode eggs in feces, is less time-consuming, separates Anoplocephala eggs from fecal debris, and decreases crystallization25. Moreover, this technique has been successfully used to detect ascarid eggs with high sensitivity26. Yet, some of these aforementioned techniques and centrifugal methods such as as the Ovassay, as opposed to the flotation protocol we propose in this study, require sample preservation in reagents such as formalin, commercial kits, sample processing under laboratory conditions, and the use of reagents such as zinc sulfate27 which are expensive and require special disposal procedures to avoid environmental toxicity.

The use of techniques that increase the sensitivity of the flotation method by adding solutions with high SpG has been favored recently. However, it must be considered that the disadvantage of these solutions is the increase in debris in the final preparation and hence, the inaccurate detection of parasite eggs. In addition, the commercial availability of materials, reagents, cost, environmental impact issues, and difficulty of use of centrifugal methods affect the selection of a flotation technique14, which can be challenging in field conditions in contrast to the protocol that we present in this work. The preparation of the flotation solutions with table salt is advantageous over the utilization of sugar because under field conditions, sugar attracts insects such as wasps and bees and preparations become sticky. Further, solutions such as phenol, which is added to sugar solutions to avoid stickiness, or ZnSO4are complex to properly discard according to environmental protection guidelines and cannot be disposed of in the field; unlike a table salt solution.

The goal of this manuscript is to demonstrate the steps to detect T. canis and Ancylostoma spp. eggs in fecal samples using an adaptation of the simple flotation technique under field and laboratory conditions. Following the protocol herewith described and using a microscope with a backup battery, the diagnosis of these canine zoonotic parasites in rural and suburban areas is possible when no laboratory equipment and infrastructure are available. The simple flotation method described in this work can provide quick results and is a non-invasive and cost-effective technique for routine screening.

Protocol

The use and care of dogs was approved by the National and Autonomous University of Mexico.

1. Collection of fecal samples

NOTE: Handle the dog with the help of a veterinarian or the owner of the animal.

  1. In the case of feral dogs (Figure 1A) or nervous animals, collect samples from the ground right after defecation or no more than 10 min later.
  2. Lubricate surgical gloves or thin-walled polyethylene bags with water or Vaseline. To collect fecal samples from the rectum, wear surgical gloves or thin-walled polyethylene bags.
  3. Collect at least 2 g of feces from each dog (Figure 1B).
  4. Identify individual fecal samples as follows: date, location (Global Positioning System [GPS] coordinates using Google Maps), assign an identification number for each animal, approximate age of the dog, gender of the dog, breed, indoor or outdoor (feral) dog.
  5. Close the bags containing the fecal sample with a tight knot. Keep the bags refrigerated (4-8 °C) if stool samples are not analyzed within 3-4 h after collection.

2. Preparation of a saturated salt solution for field diagnosis

NOTE: If access to a balance, measuring material, stoves, or gas to boil water is absent or limited, the saturated saline solution can be easily prepared using water, common table salt, a plastic 12 oz (355 mL) cup, and a 1 L soda plastic empty bottle.

  1. Wash thoroughly an empty 1 L soda bottle. Fill the bottle with 1 L of water.
  2. Fill a 12 oz plastic cup with common table salt.
  3. Transfer the salt to the soda bottle.
  4. Close tightly the soda bottle with the screw cap. Shake the solution vigorously until the salt no longer dissolves.
    NOTE: Shaking the solution in the soda bottle until complete dissolution of salt can take up to 90 min.

3. Preparation of a saturated salt solution for laboratory diagnosis

  1. Weigh 420 g of common table salt.
  2. Dissolve 420 g of salt in 1 L of water.
  3. Boil the solution until no more salt dissolves.
  4. Filter the solution to discard the undissolved salt.
  5. Check the concentration of the solution using a heavy fluid hydrometer or densitometer.
    NOTE: The ideal concentration has an SpG of 1.20 to achieve better results (Figure 2A). The floating capability of an egg is influenced by its interaction with the solution, contributing to the varying ability of eggs to float in solutions with the same specific gravity. Hence, to achieve optimal egg recovery, it is essential to consider the upper range of specific gravity in the flotation solution, ensuring it surpasses that of the targeted parasite elements6.

4. Flotation method

NOTE: If fecal samples are too dry or hard, macerate them in a mortar.

  1. Using a spoon, place approximately 3 g of feces in one plastic cup (~8.5 cm in height and ~5.5 cm in diameter).
  2. Add 1 mL of the saturated salt solution until a paste is obtained.
  3. Stir for 1 min and add 100 mL of saturated salt solution.
  4. Pass this suspension through a plastic strainer into a second plastic cup to avoid coarse particles (Figure 2B).
  5. Let the suspension stand for 15-20 min.
  6. Place an inoculation loop in a flame for 1 s to ensure it is free of eggs, cysts, or oocysts (Figure 2C).
  7. Wait for 5 s for the inoculation loop to cool down (Figure 2D).
  8. Take three drops from the surface of the suspension with the inoculation loop. Place each one of the three drops separately on one glass slide (Figure 2E-G)
    NOTE: Ensure that the drops are not in contact with each other (Figure 2H).
  9. Observe under the microscope with a 10x objective (Figure 2I). Place a coverslip if magnification is increased to 40x.
  10. When a positive result is observed in the droplet, assign a cross (+) in the laboratory logbook to record the presence of parasites in the patent period in random sites of the fecal suspension surface.

5. Interpretation of the flotation method

NOTE: Negative results are inconclusive.

  1. Perform a series of three tests with samples from 3 consecutive days to increase the sensitivity of the test.
    NOTE: Positive results indicate the presence of parasites in the patent period.

Representative Results

In this work, collection and coproparasitoscopic procedures for the identification of T. canis and Ancylostoma spp. are described. The rationale behind the adaptation of the simple fecal flotation method to detect canine helminth eggs is that this technique is cost-effective as the solutions, equipment, and materials are inexpensive. Hence, the method has a high sample handling capacity as multiple samples can be processed in a short period. Moreover, the simple fecal flotation method is easy to perform and relatively sensitive.

The sample size and choice of animals were made by convenience determined mainly by the willingness of the owner of the animals and the safety of approaching feral dogs. In the present work, the flotation method was used to assess occurrences and frequencies of Ancylostoma spp. and T. canis in dogs Canis lupus familiaris in Mexico during 4 years starting in 2017. Fecal samples from 1,638 dogs were collected and screened microscopically. A total of 1,235 dogs were positive for T. canis and Ancylostoma spp. One of the goals of employing the flotation method was to demonstrate its effectiveness in gauging the occurrence of Toxocara or Ancylostoma, but we also aimed to conduct a deeper examination of the factors that could impact their prevalence. Consequently, 185 animals with mixed infections were discarded. The number of positive samples for Ancylostoma spp. was 833 (78.95% frequency) and 222 (21.04%) for T. canis. Of the 1,050 samples, 75.5% (793 samples) were processed and read in a field setting by preparing the flotation solution with a soda bottle and using a microscope with 3 x 1.2 V AA rechargeable batteries for 4 h of continuous operation. The remaining 257 fecal samples were examined in the laboratory.

A goal of this study was to use a flotation solution and a method in a field setting to speed up and facilitate the diagnosis of two helminth parasites of dogs in areas where no laboratory infrastructure or equipment is available. The field application of the flotation method enabled us to communicate findings to 400 owners immediately after each microscopic reading to provide recommendations regarding anthelmintic treatments and preventive measures and therefore, avoid parasite spread in humans or companion animals. Likewise, the flotation technique in the field setting made it possible to examine feral dogs and provide them with an anthelmintic treatment using treats or food. Figure 3A and Figure 3B show eggs of T. canis and Ancylostoma spp., respectively, observed after the flotation solution and technique were done in a field setting. When fresh stool samples were subjected to the flotation method in a field setting, the solution prepared in a soda bottle was capable of bringing helminth eggs into flotation. Production of salt crystals and air bubbles was compared to the samples read in a laboratory setting and no difference was detected by the operator. This observation is encouraging and challenges the knowledge that the concentration of fecal samples guarantees a more accurate microscopic reading.

When the flotation solution and stool samples were processed in a laboratory setting, no difference was perceived regarding the morphology and clarity of the readings under the microscope as the same amount of debris floated concurrently in both the field and laboratory copromicroscopical detection of parasites. Figure 3A,B show T. canis and Ancylostoma spp. eggs, respectively, recovered from the 257 samples that were processed in a fully-equipped laboratory setting after transportation of samples from urban, suburban, and rural areas. These parasite eggs did not differ morphologically from the ones observed under field conditions (Figure 3C).

This protocol generated a sufficient number of findings to assess the influence of location, season of the year, gender, breed, age, and outdoor or indoor conditions in T. canis and Ancylostoma spp. prevalence. Prevalence data were compared using the ANOVA Test with a significance value of 0.05. Table 2 shows that T. canis exhibited a significantly higher frequency of distribution in temperate and dry climates compared to states characterized by warm climates, whereas there was no notable variation in the prevalence of Ancylostoma spp. across regions with predominant warm, temperate, or dry climates. Likewise, Ancylostoma spp. and T. canis are most commonly detected in summer. It was found that 60.86% of males were positive for ancylostomiasis and 37.38% for toxocariasis. Moreover, T. canis was more frequently detected in puppies less than 6 months of age; whereas Ancylostoma spp. was more diagnosed in adult dogs. Interestingly, no difference was found between dog breeds. The dog hookworm infected 65.54% of outdoor dogs; yet, T. canis was equally detected in dogs with an outdoor or indoor status.

Figure 1
Figure 1: Stool sample collection from dogs. (A) Caution must be exercised while collecting stool samples from feral dogs.When possible, fresh feces must be collected and kept in a plastic bag for submission and coproparasitoscopic analysis in the laboratory. Samples must be refrigerated at 4 °C and placed in styrofoam boxes or insulated metallic envelopes within 24 h of collection. (B) Three grams of feces are recommended as a standard; hence, 2-5 g is reasonable. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Preparation of a saturated salt solution and processing the fecal suspension for microscopy. (A) The ideal density of a saturated solution made with salt should be over 1.20. It is highly recommended to check that there are no crystals at the bottom and the solution is transparent. (B) Passing the fecal suspension through a plastic strainer removes coarse particles. (C) The inoculation loop must be exposed to a flame (a laboratory burner or a common portable lighter) to make sure that it is free of contamination. (D) Allow the inoculation loop to cool down and take three drops from different sites on the surface of the suspension. (E) After placing the first droplet on the glass slide, avoid moving the slide to reduce the spilling of the droplet. The glass slide provides a thin and transparent platform that allows for the observation of samples. (F) Place the second droplet in the middle of the glass slide. Taking a second drop from the surface of the fecal suspension increases the probability of finding eggs in a suspension where parasitic structures may have been unevenly distributed. (G) Place the third droplet on the glass slide. A third drop from the surface of the fecal suspension will allow the observation of another area where floating eggs may have concentrated. (H) The three drops are observed separately on the glass slide to detect nematode eggs from different areas of the surface of the fecal suspension. (I) Observe the sample under a microscope using the 10x objective. If the objective is changed to 40x, place a cover slip on the fecal suspension droplets. When a positive result is observed in the droplet, assign a cross (+) to mark the presence of parasites in the patent period in random sites of the fecal suspension surface. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Eggs of Toxocara canis and Ancylostoma spp. detected by light microscopy using 40x magnification. (A) Eggs of Toxocara canis detected by light microscopy using 40x magnification. The flotation method allowed the visualization of clean microscopic fields free of coarse materials (collected in a field setting). Regardless of the method of preparation of the flotation solution, eggs were visible and mainly observed without morphological alterations. (B) Eggs of Ancylostoma spp. detected by light microscopy using 40x magnification. Even though no concentration of feces was carried out (in a field setting), the simple flotation method addressed the shortcomings of morphological detection of eggs in feces when the procedure was done according to the present method. (C) Eggs of T. canis and Ancylostoma spp. detected by light microscopy using 40x magnification. Morphological features of these eggs (recovered in a laboratory setting) were clear even though no concentration of stool samples was done and did not differ from the eggs recovered in a field setting. Scale bars = 75 μm. Please click here to view a larger version of this figure.

Ancylostoma spp. Toxocara canis
Climate Warm climate 38.66a 22.52a
Dry climate 29.41a 36.03a
Temperate climate 31.93a 41.44b
Spring 23.76a 19.81a
Summer 49.81b 50.00b
Autumn 17.52a 20.27a
Winter 8.88c 9.90c
Gender Male 60.86a 62.61a
Female 39.13b 37.38b
Age 0-3 months 18.72a 51.80a
3-6 months 15.48a 32.43b
> 6 months 22.44a 10.36c
> 1 year 43.33b 5.40c
Breed Crossbred 52.78a 55.86a
Purebred 47.22a 44.14a
Housing status Indoor 34.45a 52.25a
Outdoor 65.54a 47.74b

Table 2: Prevalence (%) of Ancylostoma spp. and Toxocara canis in dogs in Mexico during 4 years according to climate, season, gender, age, breed, and housing status. Indications from the data reveal that there is no significant variation in the prevalence of Ancylostoma spp. across areas characterized by warm, temperate, or dry climates. In contrast, T. canis exhibits a notably higher prevalence in temperate and dry climates compared to warm climates. Additionally, both Ancylostoma spp. and T. canis are most frequently identified during the summer season. Analysis of the data further demonstrates that 60.86% of male dogs tested positive for ancylostomiasis, while 37.38% tested positive for toxocariasis. Furthermore, T. canis is more commonly detected in puppies aged less than 6 months, whereas Ancylostoma spp. is more prevalent in adult dogs. Interestingly, no discernible differences were observed among various dog breeds. The findings indicate that 65.54% of outdoor dogs are infected with dog hookworm, while T. canis is equally prevalent among indoor and outdoor dogs. *Mean values within a column of the factor (Climate, Season, Gender, Age, Breed and Housing) followed by the same letters are not significantly different at P < 0.05 according to ANOVA Test.

Discussion

Nematodes such as T. canis and Ancylostoma spp. can inhabit the small intestine of dogs and have the potential to be transmitted to humans. Clinical signs caused by T. canis are serious in young dogs, manifesting as poor growth, respiratory issues, or digestive tract lesions28. In adult dogs, the infection typically tends to be mild. Diagnosis relies on identifying characteristic eggs in a fecal sample. This condition is a frequent cause for prescribing anthelmintic treatment to dogs29. To understand the epidemiology of Toxocara, it is essential to recognize that the parasite can transfer from the mother to the puppy both through the placenta before birth and through suckling as a galactogenic infection shortly after whelping. Humans can contract T. canis when ingesting infective eggs, which can occur through various means such as contaminated soil or the fur of an infected dog. In this case, humans serve as paratenic hosts, and while there is no parasitic reproduction, the released larvae from the eggs can potentially cause harm, particularly in the case of children30.

Infection with the hookworm Ancylostoma spp. in dogs occurs when the animals ingest the larvae or when larvae penetrate skin. Puppies can contract the infection by consuming a paratenic host or through the lactogenic pathway31. Mature hookworms feed on blood, potentially leading to severe anemia and gastrointestinal symptoms, particularly in young dogs. Infections that occur through the skin result in dermatitis, which typically resolves about a week after the onset of the infection. Various anthelmintic medications have been employed, accompanied by supportive therapy. However, there have been reports of anthelmintic resistance. Ancylostoma has the potential to be pathogenic to humans. The transcutaneous transmission of L3 larvae can lead to a condition known as cutaneous larva migrans. These lesions generally resolve without treatment over a few months. In rare instances, the canine hookworms may migrate into the human gastrointestinal tract, inducing eosinophilic enteritis30.

The accurate, economical, and rapid diagnosis of parasitosis in dogs is essential to increase our epidemiological knowledge of dog helminthiases. For this reason, we adapted a protocol of the simple flotation technique to be used in a field setting to detect T. canis and Ancylostoma spp. A relevant issue faced in the current protocol was the collection of samples from feral dogs, as these animals are usually hard to handle. Therefore, sampling was done by observing and following the dogs until they defecated, making the sampling method time-consuming, challenging, and somehow dangerous. According to our experience, we strongly recommend assessing parasitological studies in shelter dogs.

Regarding the zoonotic helminth egg recovery, qualitative coproscopy continues to be the methodology commonly used in the parasitology laboratory for diagnostic purposes1. Previous studies have found that the flotation technique using a saturated salt solution is more reliable than direct microscopy. For example, Dryden and collaborators determined a sensitivity of 95.15% for Ancylostoma spp. eggs in 206 fecal samples processed by the flotation technique and 27.18% by direct microscopy32. It is accepted that the density of the flotation solution is a critical factor for the flotation method14,20,23. In solutions that do not reach that density, the parasitic forms take longer to float or never float. When a flotation solution is oversaturated, it crystallizes in a couple of minutes and even earlier if a coverslip is placed20.

The choice of the flotation solution in this protocol was based on ease of preparation, convenience, cost, and environmental impact. Even though the sugar solution takes longer to crystallize, formalin or phenol should be added as a preservative and decrease the sticky or gluey sensation; which is an inconvenience under the field conditions that prevailed in this study. Moreover, the disposal of chemical reagents is a complex issue due to environmental regulations. Data in this study demonstrated that the saturated solution prepared with common table salt under field conditions by using a soda bottle due to the lack of availability of laboratory equipment to prepare and boil a flotation solution, was as effective as the one prepared in a laboratory setting. The only perceptible difference is the longer time it takes to shake the soda bottle until a transparent solution can be seen. Microscopic observations were equally clean for the sample prepared using both methods to prepare the saturated solution. It is important to perform more studies in the future with a larger number of samples to confirm the aforementioned statement. Yet, it is time-consuming and needs the physical effort of shaking the solution inside the bottle for hours. Another critical step of the protocol consists of letting the fecal suspension stand for at least 15 min to allow the eggs to float and concentrate on the surface. However, the longer the resting time, the more debris will float and after a few hours, the eggs will deform or break. Considering that sensitivity and specificity of coproparasitoscopic methods are low and that accuracy of these methods as diagnostic tests depends on the skills of the user, in the current study, trained and experienced parasitologists performed the flotation technique and identified the parasites.

This study, to our knowledge, is the first to estimate the prevalence of Ancylostoma spp. and T. canis in dogs in Mexico using fecal samples examined with the simple flotation method in a field setting. It was herein demonstrated that the flotation technique we used, despite the decades it has been in use, is still a highly effective, economical, and rapid diagnostic test14. Even so, there are coproparasitoscopic techniques with higher sensitivity and specificity than the one we used for the diagnosis of dog helminths; such as the sedimentation concentration, as is the case of the Faust technique. Furthermore, it is important to note that Faust, which is a concentration method, has also been shown to be more effective in the detection of protozoa such as Giardia33. Nonetheless, the coproparasitoscopic method we chose for this study had several advantages such as cost. The Faust technique is more expensive than flotation with saline-saturated solution because it requires additional and expensive equipment such as a centrifuge. Other concentration methods such as those using reagents such as Zinc sulfate are more expensive than common salt and need more time due to the number of the required washes in the centrifuge. Therefore, the Faust technique is limited for its use in field work where no access to a centrifuge is common.

Most importantly, a limitation of the simple flotation technique is that it is not sensitive enough to detect Cryptosporidium oocysts, Giardia cysts, or Trichuris eggs. Techniques using centrifugation procedures to concentrate parasite forms should hence be used if the diagnosis of these species is intended. It holds significance to highlight that the positivity rates of the genera reported in this study should not be regarded as conclusive because a convenience method was used as a sampling strategy. It is important to note that there might be situations, such as in shelters with challenging animal management, where it becomes crucial to identify helminth parasites in dogs. As a result, this protocol may need adjustments in the future to accommodate the collection of combined samples.

As a weak point, it should be noted that the simple flotation method and the preparation of the saturated solution of NaCl were developed for a specific purpose-to examine dog feces under mostly field conditions-but the sensitivity of the simple flotation procedure has generally been considered to be low. Therefore, to detect cysts or eggs of heavier parasite species, the use of a concentration technique is strongly suggested. Given the low sensitivity of the simple flotation method, regardless if it is performed in a field or laboratory setting, it was reasonable to doubt if this technique would be useful as a method of choice for epidemiological studies of canine helminth parasites. Nonetheless, the high Ancylostoma spp. prevalence in dogs in this study and the detection of T. canis eggs indicate that the simple flotation method, whether performed in a field or laboratory setting, is useful to provide evidence that might support further epidemiological studies and propose control and preventive measures to reduce the infection of dogs and humans with these nematodes.

Using the simple flotation method, we analyzed the factors affecting T. canis and Ancylostoma spp. egg recovery from canine feces. T. canis findings were in agreement with the latest research that indicates that parasites are notably more prevalent in temperate and tropical regions. This aligns with prior studies that have investigated how environmental factors, particularly temperature and rainfall, influence the ecological range of parasite species34,35,36. The present results showed that both helminths infect males in a significantly higher proportion. It is reasonable to speculate that this finding was because males are immunologically less reactive to infections37,38. Age influences the prevalence of T. canis as puppies were more affected by this parasite. This finding aligns with earlier research indicating that this parasite tends to impact puppies to a greater extent, while adults develop increased resistance, resulting in a lower likelihood of infection6,39. Both crossbred and purebred dogs showed comparable prevalences of Toxocara canis and Ancylostoma spp. However, feral dogs exhibited a higher prevalence of Ancylostoma spp. compared to dogs residing in domestic settings. This finding is in disagreement with previous reports that did not detect differences between dog breeds6. It can be concluded that future applications of the preparation of the saturated saline solution and the simple flotation method used in this protocol can be conducted for routine fecal examinations under field or laboratory conditions.

Açıklamalar

The authors have nothing to disclose.

Acknowledgements

The authors are grateful to the Dirección General de Asuntos del Personal Académico of the Universidad Nacional Autónoma de México for providing the financial resources through grant PAPIIT IN218720 and to Dr. Claudia Mendoza for granting the requested extension. This work is dedicated to my lovely Nicole, who passed away in 2019. You will always live in my heart.

Materials

3 x 1.2 V AA rechargeable batteries Energizer Sold in retail stores
Bunsen burner Viresa FER-M224
Disposable 12-oz glass cup Uline Mexico S-22275 Sold in retail stores
Glass slides Velab, Mexico VEP-P20
Inoculating loop VelaQuin, Mexico CRM-5010PH 
Light Microscope VelaQuin, Mexico VE-B2
Lighter Bic J25 Sold in retail stores
One plastic cup (12 oz) Amazon ASIN B08C2CRHSH Can be any kitchen plastic reuseable cup
Plastic cups  (size of a dice or urine sample cup) diameter 5.5 cm and height 8.5 cm, two cups Amazon Layhit-Containers-ZYHD192919 Can be any kitchen plastic reuseable cup
Plastic strainer 10 cm Ecko ASIN B00TUAAVWI Can be any kitchen plastic strainer
Soda bottle Coca-Cola 1-liter Sold in retail stores
Spoon Amazon Basics ASIN B00TUAAVWI Can be any kitchen spoon
Table salt La Fina Sold in retail stores
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Referanslar

  1. Duncan, K. T., Koons, N. R., Litherland, M. A., Little, S. E., Nagamori, Y. Prevalence of intestinal parasites in fecal samples and estimation of parasite contamination from dog parks in central Oklahoma. Veterinary Parasitology Regional Studies and Reports. 19, 100362 (2020).
  2. Kisiel, L. M., et al. Owned dog ecology and demography in Villa de Tezontepec, Hidalgo, Mexico. Preventive Veterinary Medicine. 135, 37-46 (2016).
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  7. Cortez-Aguirre, G. R., Jiménez-Coello, M., Gutiérrez-Blanco, E., Ortega-Pacheco, A. Stray dog population in a city of Southern Mexico and its impact on the contamination of public areas. Veterinary Medicine International. 2018, 1-6 (2018).
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  12. Pastor-Santiago, J. A., Flisser, A., Chávez-López, S., Guzmán-Bracho, C., Olivo-Díaz, A. American visceral leishmaniasis in Chiapas, Mexico. The American Journal of Tropical Medicine and Hygiene. 86 (1), 108-114 (2012).
  13. Trasviña-Muñoz, E., et al. Detection of intestinal parasites in stray dogs from a farming and cattle region of Northwestern Wexico. Pathogens. 9 (7), (2020).
  14. Ballweber, L. R., Beugnet, F., Marchiondo, A. A., Payne, P. A. American Association of Veterinary Parasitologists’ review of veterinary fecal flotation methods and factors influencing their accuracy and use–is there really one best technique. Veterinary Parasitology. 204 (1-2), 73-80 (2014).
  15. Nielsen, M. K. What makes a good fecal egg count technique. Veterinary Parasitology. 296, 109509 (2021).
  16. Gordon, H. M., Whitlock, H. V. A new technique for counting nematode eggs in sheep faeces. Journal of the Council for Scientific and Industrial Research. 12 (1), 50-52 (1939).
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  18. ESCCAP. Worm control in dogs and cats. Guideline 01. European Scientific Counsel Companion Animal Parasites. , (2017).
  19. Cringoli, G., et al. Ancylostoma caninum: Calibration and comparison of diagnostic accuracy of flotation in tube, McMaster and FLOTAC in faecal samples of dogs. Experimental Parasitology. 128 (1), 32-37 (2011).
  20. Figueroa, C. J. A. Examen coproparasitoscópico. In: Técnicas para el diagnóstico de parásitos con importancia en salud pública y veterinaria, Consejo Técnico Consultivo Nacional de Sanidad Animal. AMPAVE-CONASA. , 83-105 (2015).
  21. Dryden, M. W., Payne, P. A., Ridley, R., Smith, V. Comparison of common fecal flotation techniques for the recovery of parasite eggs and oocysts. Veterinary Therapeutics. 6 (1), 15-28 (2005).
  22. Adolph, C., et al. Diagnostic strategies to reveal covert infections with intestinal helminths in dogs. Veterinary Parasitology. 247, 108-112 (2017).
  23. Cringoli, G., Rinaldi, L., Maurelli, M. P., Utzinger, J. FLOTAC: New multivalent techniques for qualitative and quantitative copromicroscopic diagnosis of parasites in animals and humans. Nature Protocols. 5 (3), 503-515 (2010).
  24. Katagiri, S., Oliveira-Sequeira, T. C. Comparison of three concentration methods for the recovery of canine intestinal parasites from stool samples. Experimental Parasitology. 126 (2), 214-216 (2010).
  25. Rehbein, S., Lindner, T., Visser, M., Winter, R. Evaluation of a double centrifugation technique for the detection of Anoplocephala eggs in horse faeces. Journal of Helminthology. 85 (4), 409-414 (2011).
  26. Liccioli, S., et al. Sensitivity of double centrifugation sugar fecal flotation for detecting intestinal helminths in coyotes (Canis latrans). Journal of Wildlife Diseases. 48 (3), 717-723 (2012).
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  28. Barutzki, D., Schaper, R. Endoparasites in dogs and cats in Germany 1999-2002. Parasitology Research. 90, S148-S150 (2003).
  29. Carlin, E. P., Tyungu, D. L. Toxocara: Protecting pets and improving the lives of people. Advances in Parasitology. 109, 3-16 (2020).
  30. Saari, S., NaReaho, A., Nikander, S. . Canine parasites and parasitic diseases. , (2019).
  31. Bowman, D. D., Montgomery, S. P., Zajac, A. M., Eberhard, M. L., Kazacos, K. R. Hookworms of dogs and cats as agents of cutaneous larva migrans. Trends in Parasitology. 26 (4), 162-167 (2010).
  32. Dryden, M. W., Payne, P. A., Ridley, R., Smith, V. Comparison of common fecal flotation techniques for the recovery of parasite eggs and oocysts. Veterinary Therapeutics. 6 (1), 15-28 (2005).
  33. Barutzki, D., Schaper, R. Results of parasitological examinations of faecal samples from cats and dogs in Germany between 2003 and 2010. Parasitology Research. 109, S45-S60 (2011).
  34. Novobilský, A., Novák, J., Björkman, C., Höglund, J. Impact of meteorological and environmental factors on the spatial distribution of Fasciola hepatica in beef cattle herds in Sweden. BMC Veterinary Research. 11, 128 (2015).
  35. Pickles, R. S., Thornton, D., Feldman, R., Marques, A., Murray, D. L. Predicting shifts in parasite distribution with climate change: A multitrophic level approach. Global Change Biology. 19 (9), 2645-2654 (2013).
  36. Pérez-Rodríguez, A., De La Hera, I., Fernández-González, S., Pérez-Tris, J. Global warming will reshuffle the areas of high prevalence and richness of three genera of avian blood parasites. Global Chaneg Biology. 20 (8), 2406-2416 (2014).
  37. Nava-Castro, K., Hernández-Bello, R., Muñiz-Hernández, S., Camacho-Arroyo, I., Morales-Montor, J. Sex steroids, immune system, and parasitic infections: Facts and hypotheses. Annals Of The New York Academy Of Sciences. 1262, 16-26 (2012).
  38. Morales-Montor, J., Escobedo, G., Vargas-Villavicencio, J. A., Larralde, C. The neuroimmunoendocrine network in the complex host-parasite relationship during murine cysticercosis. Current Topics in Medicinal Chemistry. 8 (5), 400-407 (2008).
  39. Olave-Leyva, J., et al. Prevalencia de helmintos gastrointestinales en perros procedentes del servicio de salud de Tulancingo, Hidalgo. Abanico Veterinario. 9 (1), 1-10 (2019).

Etiketler

Fecal FlotationZoonotic NematodesToxocara CanisAncylostoma Spp.Field DiagnosisLaboratory DiagnosisCanine ParasitesDewormingEpidemiologyCoproparasitoscopic DetectionCryptosporidiumGiardiaTrichuris

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Segura, J., Alcala-Canto, Y., Figueroa, A., Del Rio, V., Salgado-Maldonado, G. A Simple Fecal Flotation Method for Diagnosing Zoonotic Nematodes Under Field and Laboratory Conditions. J. Vis. Exp. (202), e66110, doi:10.3791/66110 (2023).
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