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Guide for the Care and Use of Laboratory Animals

A respected resource for decades, the Guide for the Care and Use of Laboratory Animals has been revised by a committee of experts, based on input from scientists and the public. The Guide incorporates recent research on commonly used species, including farm animals, and includes extensive references. It is organized around major components of animal use:

• Institutional policies and responsibilities. The committee discusses areas that require policy attention: the role and function of the Institutional Animal Care and Use Committee, protocols for animal care and use, occupational health and safety, personnel qualifications, and other areas.
• Animal environment, husbandry, and management. The committee offers guidelines on how to design and run a management program, addressing environment, nutrition, sanitation, behavioral and social issues, genetics, nomenclature, and more.
• Veterinary care. The committee discusses animal procurement and transportation, disease and preventive medicine, and surgery. The Guide addresses pain recognition and relief and issues surrounding euthanasia.
• Physical plant. The committee identifies design and construction issues, providing guidelines for animal-room doors, drainage, noise control, surgery, and other areas.

The Guide for the Care and Use of Laboratory Animals provides a framework for the judgments required in the management of animal facilities—a resource of proven value, now updated and expanded. This revision will be important to researchers, animal care technicians, facilities managers, administrators at research institutions, policymakers involved in research issues, and animal welfare advocates.

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At the Multilateral Consultations held in 1992, 1993 and 1997, the Parties adopted Resolutions aiming at facilitating and harmonising the implementation of certain provisions of the Convention.

For their work, the Parties have invited non-member States of the Council of Europe and cooperated very closely with non-governmental organisations representing the fields concerned: scientists, veterinarians, laboratory animal breeders, animal protection associations, specialists in animal science, representatives of the pharmaceutical industry etc. who participated as observers in the meetings.

In June 1998, a Protocol of Amendment (ETS No. 170) providing for a simplified procedure for the amendment to the technical appendices to the Convention was opened for signature.

In 1998, the Parties to the Convention undertook a revision of Appendix A to the Convention, on the accommodation and care of animals. The Working Party completed the revision of the Appendix at its 8th meeting, from 22 to 24 September 2004, and submitted it to the Multilateral Consultation of the Parties for adoption.

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The Appendix entered into force twelve months after its adoption, i.e. on 15 June 2007, as required by Article 31 of the Convention for the Protection of Vertebrate Animals used for Experimental and other Scientific Purposes.

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The purpose of the Guide for the Care and Use of Laboratory Animals (the Guide ), as expressed in the charge to the Committee for the Update of the Guide , is to assist institutions in caring for and using animals in ways judged to be scientifically, technically, and humanely appropriate. The Guide is also intended to assist investigators in fulfilling their obligation to plan and conduct animal experiments in accord with the highest scientific, humane, and ethical principles. Recommendations in the Guide are based on published data, scientific principles, expert opinion, and experience with methods and practices that have proved to be consistent with both high-quality research and humane animal care and use. These recommendations should be used as a foundation for the development of a comprehensive animal care and use program, recognizing that the concept and application of performance standards, in accordance with goals, outcomes, and considerations defined in the Guide , is essential to this process.

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Management of Laboratory Animals

• Sanitation |
• Vermin Control |

Consistently delivered quality programs of husbandry and veterinary care provide the foundation that enables valid scientific research. For proper management of research animals, the animal care and research staff must be responsible, sensitive to the animals’ health and well-being, well trained in the humane care and use of laboratory animals, highly motivated, experienced, and diligent in performing their duties and responsibilities. Standard operating procedures are valuable, and training and supervision are essential to assure a consistently applied and uniformly high level of animal care. Within research facilities, environmental conditions must be carefully controlled so that, along with conscientiously applied programs of animal care and use, the best possible conditions for conducting research are provided.

The Guide remains the primary reference for information on basic principles and standards for laboratory animal management. Laboratory rodents that are disease- and pathogen-free and that do not possess antibodies indicative of past infection are readily available from commercial vendors. Procuring such animals from high-quality sources, transporting them in filtered shipping containers, and maintaining them in facilities with both physical and procedural barriers to the introduction of infectious agents are effective measures to prevent disease within a colony that may confound or ruin experiments.

For certain studies, particularly those associated with immunity, it has become clear that animals protected from exposure to a broad array of organisms have an immune repertoire insufficiently developed to be suitable. This requires adaptable management programs to maintain and segregate subjects across an expanse of health profiles from axenic and gnotobiotic to those in close proximity or intentionally exposed to and potentially harboring a medley of otherwise undesirable pathogens.

Although there are colonies of some species of primates that are free of most agents that cause infectious disease in these species, many primates used are of feral or wild origin. For this reason, appropriate quarantine, isolation, and conditioning programs are critical, in addition to the program followed in the importers’ facilities.

Housing Considerations for Laboratory Animals

Cages, pens, or runs should provide adequate space to allow for normal physiologic needs, permit postural adjustments, and meet requirements for species-specific behavior. When possible, compatible groups of social animals should be housed together. Primary enclosures should be constructed of durable materials, easily cleaned and sanitized, and designed for comfort and safety. Static microisolation (filter-top) cages and individually ventilated caging (IVC) systems impede cage-to-cage transmission of infectious agents. However, infection can be transmitted horizontally or vertically from parents to progeny in breeding colonies; naive mice introduced for cross-breeding and back-crossing can perpetuate infection; and experimental mice potentially can be exposed to pathogens via a contaminated environment, shared watering valves, research devices, or when taken to laboratories. Individual ventilation of cages serves to delay deterioration of the environment within the cage and maintain a more consistent and wholesome microenvironment; it also saves space in the facility and can be engineered to minimize odors, allergens, dust, and heat exhausted into the macroenvironment.

Federal law in the USA requires that laboratory dogs have an opportunity to exercise regularly and have sensory contact with other dogs unless restricted by experimental or behavioral considerations. Housing for nonhuman primates must provide social and environmental enrichment to promote their psychological well-being compatible with the experimental and practical constraints of the housing situation. Successful enrichment strategies for nonhuman primates have included pair or group housing; variation in the dietary content and method of presentation; diversification of the internal cage environment with ancillary equipment (eg, perches, swings, or ladders); provision of devices to enhance visual, auditory, or tactile stimulation; and participation in challenging, nonaversive behavioral laboratory studies. Efforts to extend and adapt environmental enrichment practices to other laboratory animal species warrant consideration.

Temperature, relative humidity, ventilation rates, lighting conditions (spectrum, intensity, and photoperiod), gaseous pollutants (eg, ammonia), and noise should be carefully controlled at all times and monitored as appropriate. Unstable environmental conditions can have a profound effect on the comfort, well-being, and metabolism of animals and therefore on the quality of experimental data derived.

In general, air temperature should take into consideration the comfort ranges where human workers perform best of 64–73°F (18–22°C) and be set at +/- 2°F of set point. Ranges to take into consideration for optimal well-being of adapted animals are given in the accompanying table. Emerging evidence suggests that mice, whether adults, neonates, hirsute or not, in particular, have been kept traditionally under cold-stress conditions, especially in IVC, to the possible detriment of colony production and many types of experiments, especially those reliant on the immune system or associated with metabolism or physiology. Experience suggests, however, that mice provided an adequate energy diet and housed socially can compensate via nest building and huddling. Relative humidity should be maintained at 30%–70% for most species and preferably within 10% of the set point.

Preferred Temperature Ranges for Commonly Used Laboratory Animals

Ventilation rates should be 10–15 fresh air changes/hour but can be reduced in some situations to save energy (eg, rodent IVC with rack effluent directly connected to the room exhaust). Air recirculation is discouraged unless treated to remove particulate and gaseous contaminants.

Evenly distributed and sufficiently intense lighting promotes animal well-being, circadian rhythmicity, and allows personnel to observe the animals and perform all husbandry and sanitation duties safely and effectively. Diurnal or day-night cycles, as determined by species’ requirements, should be controlled by automatic timers to maintain circadian and neuroendocrine regulation. The introduction of LED for lighting, although offering many advantages known to consumers, influences circadian rhythms differently than cool, white fluorescent lighting traditionally in use and represents arguably the most momentous environmental change influencing experiments since the removal of windows and exposure to seasonal daylight from animal research facilities decades ago.

The microenvironment within certain types of caging may be very different from that of the macroenvironment of the room. Carefully conducted additional research is needed to more precisely define the optimal environmental and social conditions for each species or group of species at the cage level. Incomplete description of the environment in association with published animal experiments has been a longstanding flaw contributing to the contemporary crisis of unreliability in reproducing some animal experiment results over time and across laboratories.

Bedding Considerations for Laboratory Animals

Animal bedding is a controllable environmental factor that can influence experimental data and animal well-being. Bedding materials should be nonirritating, absorbent, free of chemical contamination and pathogens, and unpalatable, to discourage digestion. Adequate quantities are vital to keep animals dry and clean between changes of bedding or caging.

Rodents are the order of mammals maintained most commonly on contact bedding, of which ground corncobs, hardwood chips, recycled paper, heat-treated softwood shavings, or virgin cellulose are most commonly used. Untreated softwoods are not recommended because they contain volatile oils that may alter hepatic enzyme systems and affect certain kinds of research. This variation in product type has the potential to introduce subtle variability to reproduction, behavior, physiology, mucosal immunity, dietary studies, tactile perception, the microbiome, and environmental microorganisms.

Bedding materials also vary in their vitamin C and phytoestrogen content and ability to suppress ammonia, the byproduct of fecal bacterial urease catalysis of urea and the most abundant, noxious gas produced in the microenvironment. Depending on research requirements, bedding may be sterilized by autoclaving or irradiation before use or may be used as is. Mice, the species maintained in highest numbers on contact bedding, prefer softer materials and those in volumes that allow burrowing and nesting, supplemented with enrichment facilitating these and other normal behaviors. In light of these facts, those who care for and use animals in research should be cognizant that the wide variety in bedding materials may have equally broad effects upon behavior and biological processes that may influence experimental outcomes.

Feeding of Laboratory Animals

Feed should be of adequate quantity, palatable, free of contaminants, nutritionally adequate, easily accessible, and provided using means that meet behavioral needs according to specific species requirements. Feeds specifically manufactured for research animal use are preferred, because they are more likely to be uniformly constituted, free of contaminants, of known shelf life, and mill dated. Feed should be manufactured, transported, stored, and used in ways that minimize its deterioration, contamination, or infestation. Diets for laboratory animals generally have a 9-month shelf life if stored at or below 70°F (21°C) and 50% relative humidity. Owing to use of a stabilized type, this is the case even for diets fortified with vitamin C.

Most small animals consume food in relation to their energy requirements as influenced by the environment and dictated by their genotype and are fed ad lib; to prevent obesity and reduce wastage, rabbits, laboratory carnivores, swine, aquatic amphibians, and primates may be restricted to measured quantities of feed each day. As a general rule, laboratory animals minimally consume 4%–6% of their body weight in food daily. In addition to commercially prepared and usually pelleted natural ingredient diets of varying specification (eg, quality control and assurance of ingredients), semisynthetic or completely synthetic diets and all-liquid preparations can be formulated for use in certain kinds of research. Autoclavable or irradiated diets are available for rodents and can be used when sterilization of feed is desired.

Water Requirements of Laboratory Animals

For terrestrial animals, potable, uncontaminated water must be provided in adequate quantities to meet specific species requirements. Water entering an animal research facility will typically be supplied from a local, domestic source (although sometimes from wells), meeting general standards appropriate for human consumption. This water, however, may still be subject to considerable variation depending upon a number of factors, including geographic locale, the proximity to industrial, urban, or agricultural settings, and municipal treatment approaches. Drinking water consequently is a variable that can profoundly affect research.

Prior to presentation for consumption, water entering the animal research facility should undergo sediment and carbon filtration, water softening, and additional purification, such as deionization or reverse osmosis filtration, prior to storage in reservoirs of capacity with multiple days’ supply in reserve and distribution for animal consumption. Quality assurance programs that monitor pH, hardness, chemical content, and microbial load are recommended. Acidified, chlorinated, or sterile water may be required under certain experimental or husbandry conditions.

Water is usually provided ad lib via bottles, plastic pouches, via automated delivery, and, in certain applications, nonwetting, sterile gels. Particularly in the housing of rodents, an automated water supply enhances the advantages of ventilated caging systems and reduces operational costs/expenses, increases safety for animal care technicians, saves labor, reduces disruptions of the mice by caretakers, and provides consistently high water quality. The drawback of the use of automated drinking water supply for rodents is the risk of hypothermia, drowning, or dehydration of cage inhabitants as a consequence of failure of the in-cage water delivery valve. These systems do not operate automatically and require regular oversight of function and periodic preventive maintenance, including full distribution system sanitization to prevent biofilm formation.

Water quality is the most important environmental variable for aquatic species and a key determinant of health. Inadequate water quality or fluctuations of water temperature are physiologic stressors that impact the intake, digestion, and use of food; alter the immune system; and predispose to opportunistic infection. Water for aquatic vertebrates should be free of nitrite, ammonia, and chlorine, with total coliform counts not exceeding 200/mL. The pH should be 6.5–8.5. Although aquatic amphibians may be maintained in small containers of standing water, water recirculation with biologic filtration and periodic partial replenishment with fresh water, just as with fish, are helpful in suppressing bacterial counts and preventing the build-up of toxic chemicals.

Sanitation Considerations for Laboratory Animals

A uniformly high level of animal enclosure and facility sanitation is mandatory to ensure that animals are clean and dry, air quality is adequate (without using masking agents), and primary enclosure surfaces and accessories are clean. Housing rooms, ancillary support spaces, and primary enclosures should be cleaned and sanitized as often as necessary to keep them free of dirt, debris, and potentially harmful contamination. For rodents in solid-bottom cages, usually 1–3 changes per week will suffice; for rodents, rabbits, and nonhuman primates in suspended cages over excreta pans and for mice in ventilated caging systems, cage changes every other week should be adequate. For larger animals, excreta and soiled bedding should be removed daily, and primary enclosures cleaned and sanitized daily, or at minimum every other week.

Water bottles and other watering or feeding devices should be cleaned and sanitized at least weekly. Automated watering devices on cages, racks, or in rooms should be designed and programmed to flush continually or regularly or they should be manually drained, rinsed, and sanitized at regular, frequent intervals.

Heating cages and other equipment to 180°F (82.2°C) or using appropriate chemical disinfection (eg, hypochlorite solutions) kills nonsporeforming pathogenic bacteria and viruses. All caging and other equipment should be rinsed thoroughly after treatment with detergents or disinfectants. The effectiveness of the programs of sanitation should be evaluated regularly using appropriate microbiologic, organic material detection systems or other means.

Vermin Control Considerations for Laboratory Animals

Professionally directed programs to prevent, identify, and eradicate or control insects or escaped, feral, or wild rodents must be instituted, regularly scheduled, and consistently documented. The use of pesticides should be as a last resort and generally be confined to areas not used for animals or for storage of feed or bedding. If these agents are used in proximity to animals or their food and bedding, researchers should be informed of the use. Relatively inert substances, such as silica aerogel or boric acid powder, are recommended and are useful for control of crawling insects, eg, cockroaches.

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Care and Handling of Laboratory Animals

• First Online: 16 November 2022

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Care of laboratory animals is one of the essential parts of biomedical research. The laboratory animals such as a mouse, rat, guinea pig, rabbit, hamster, gerbils, ferret, dog, and cat should be maintained in a hygienic environment provided with good ventilation, day/night cycle, and humidity condition. The living environment of the experimental animals should be maintained uniformly throughout their lifespan to minimize stress and anxiety. Handling is an important part of the maintenance of laboratory animals. Inappropriate handling of laboratory animals makes them become stressed and anxious. Importantly, the researcher or animal handler should have appropriate training and may attend the courses related to animal welfare to eliminate or minimize stress. In this chapter, care and handling of laboratory animals are discussed.

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Care and Maintenance of Laboratory Animals

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Parasuraman, S. (2022). Care and Handling of Laboratory Animals. In: Lakshmanan, M., Shewade, D.G., Raj, G.M. (eds) Introduction to Basics of Pharmacology and Toxicology. Springer, Singapore. https://doi.org/10.1007/978-981-19-5343-9_3

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• Published: 29 September 2004

Use of animals in experimental research: an ethical dilemma?

• V Baumans 1 , 2  

Gene Therapy volume  11 ,  pages S64–S66 ( 2004 ) Cite this article

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Mankind has been using animals already for a long time for food, for transport and as companion. The use of animals in experimental research parallels the development of medicine, which had its roots in ancient Greece (Aristotle, Hippocrate). With the Cartesian philosophy in the 17th century, experiments on animals could be performed without great moral problems. The discovery of anaesthetics and Darwin's publication on the Origin of Species, defending the biological similarities between man and animal, contributed to the increase of animal experimentation. The increasing demand for high standard animal models together with a critical view on the use of animals led to the development of Laboratory Animal Science in the 1950s with Russell and Burch's three R's of Replacement, Reduction and Refinement as guiding principles, a field that can be defined as a multidisciplinary branch of science, contributing to the quality of animal experiments and to the welfare of laboratory animals. The increased interest in and concern about animal welfare issues led to legislative regulations in many countries and the establishment of animal ethics committees.

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Van Zutphen LFM . History of animal use. In: Van Zutphen LFM, Baumans V, Beynen AC (eds). Principles of Laboratory Animal Science . Elsevier: Amsterdam, 2001, pp 2–5.

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Russell WMS, Burch RL . The Principles of Humane Experimental Technique . Methuen: London, 1959, Reprinted by UFAW, 1992: 8 Hamilton Close, South Mimms, Potters Bar, Herts EN6 3QD England.

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Baumans, V. Use of animals in experimental research: an ethical dilemma?. Gene Ther 11 (Suppl 1), S64–S66 (2004). https://doi.org/10.1038/sj.gt.3302371

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Published : 29 September 2004

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• Helminthologia
• v.56(2); 2019 Jun

Intestinal Parasites of Pets and Other House-kept Animals in Moscow

O. p. kurnosova.

1 All-Russian Scientific Research Institute of Fundamental and Applied Parasitology of Animals and Plants named after K.I. Skryabin - Branch of the Federal State Budget Scientific Institution, «Federal Scientific Center - All- Russian Scientific Research Institute of Experimental Veterinary Medicine named after K.I. Skryabin and Y.R. Kovalenko the Russian Academy of Sciences», 28, Bolshaya Cheremushkinskaya street, Moscow, Russia, 117218

M. V. Arisov

I. m. odoyevskaya.

A study screening pet animals (dogs, cats, chinchillas, ferrets, guinea pigs, rabbits, primates, reptiles, and hedgehogs) within Moscow city limits for intestinal parasitic diseases has been conducted over a period of 6 years. According to the study, parasitic infections caused by intestinal protozoa are found in pet animals more frequently than by intestinal helminths. Although dogs and cats exhibit the highest level of diversity of intestinal parasite species, in the group of exotic animals, helminth infection are found much less frequently and parasitic fauna is represented mostly by intestinal protozoa with a high percentage of mixed infection. The most widespread helminth infeсtion of dogs and cats is toxocarosis (respectively 2.5 and 5.7 %) and the most widespread protozoan infection is Giardia sp. (9.8 and 4.6 %). Giardia sp. was found in 47.4 % of chinchillas, Cryptosporidium sp. was more frequently found in ferrets (6.55 %), protozoa from the family Trichomonadida was found in guinea pigs (9 %), Eimeria sp. in rabbits (13.9 %), Acanthocephal a in primates (15.7 %), and eggs from the genera Oxyurida (59 %), along with protozoa from the family Trichomonadida, in reptiles. Capillaria sp. was most prevalent in hedgehogs (33.4 %). Acanthocephala eggs, as well as protozoa from the Giardia and Entamoeba genera, were more frequently found in primates. Parasites common to animals and humans, which may become a source of infection for the latter under certain conditions, have been identified in pet animals.

Introduction

Moscow is home to a huge number of pet animals, and most of them are cats and dogs. Ferrets, guinea pigs, chinchillas, and rabbits are also often kept in urban apartments. The number of owners who keep hedgehogs, rats, reptiles, and birds is smaller. Small animal enclosures in large shopping malls that often hold squirrels and primates have recently become very popular. Also popular are petting zoos with guinea pigs, rabbits, and mini pigs. Animals have become integral to the life of people living in major metropolitan areas. In most cases, they are full-fl edged family members.

Constant veterinary supervision of animal health status, various kinds of preventative activities conducted on a constant basis, the owner’s knowledge of animal diseases and their regular visiting city’s veterinary clinics contribute to the reduction of some infectious diseases of the animals in the city and improve their quality of life in the event of chronic conditions. Without a doubt, however, certain diseases continue to be a matter of urgency despite the preventative measures that are being taken in a regular manner. Intestinal parasitic diseases caused by various species of helminths and protozoa represent a special case among these pet animal diseases. For this group of diseases, urban areas are an unique environment that creates preconditions for circulating pet animal’s infections due to widespread importation of animals from other countries and introduction of previously unseen parasite types ( Kurnosova, 2009 ). The high concentration of pet dogs, presence of stray animals, use of common walking areas, and inadequate feces collection within city limits tend to maintain and transmit the source of infection to other animals ( Erofeeva & Doronina, 2017 ). Many parasitic diseases, especially helminthiases, are low-key infections that are subclinical. In such cases, pet animals become a source of infection for an extended period of time. The course and duration of the desease depends on many factors, such as the type and intensity level of the infestation, the age and immune system status of the animal, and the presence of any concomitant underlying diseases. In general, intestinal parasitic diseases can seriously harm the health of the animal ( Kurnosova, 2009 ; Panova, 2011 ; Stepanov, 2014).

There are parasitic diseases common to animals and humans that pose a serious threat to the latter (toxocarosis, alveolar echinococcosis, cystic echinococcosis, and toxoplasmosis) ( Malysheva et al., 2013 ; Shcheveleva et al., 2016 ). Due to the biological features of some parasitic diseases, the risk of direct infection from infected animal to human is very low, because some helminth or protozoan types need a certain environment residence time, temperature, and humidity to be able to infect. There is, however, a group of so-called “contact” parasitic diseases whose infecting principle is introduced into the environment while already primed for infestation and can be transmitted to humans under certain conditions (dwarf tapeworms, enterobiasis, giardiosis, cryptosporidiosis, blastocystosis, many species of amoebas) ( Sergiev et al., 2006 ). If personal hygiene is inadequate, shelter staff, pet owners, and young children are at risk.

Many major cities are regularly screening pet animals and stray animals for parasitic diseases: St.Petersburg ( Prozorov, 1999 ); Novosibirsk ( Zubareva, 2001 , Konyaev & Borcova, 2015 ); Kostroma ( Mikhina, 2008 ); Moscow ( Kurnosova, 2009 ; Panova, 2011 ; Uspensky et al., 2011 ; Loshchinin et al., 2015 ; Kurnosova & Odoyevskaya, 2017); London (Upjonh & Cobb, 2010); Italy, Netherlands, United Kingdon (Feng & Xiao, 2011), Calgary ( Joffe & Van Niekerk, 2011 ); Austria, France, Italy, Portugal ( Beugnet & Bourdeau, 2014 ; Banerh & Thamsbord, 2016 ; Pereira & Martins, 2016 ). Studies of this type are of particular value if the duration of observation is sufficiently long. In addition to identifying the species composition of the pet animal parasites circulating throughout the city, such a study makes it possible to register new species, take timely remedial and preventative measures, and predict new developments related to the most widespread infections. This publication provides observational data obtained in a 6-year study of disease prevalence in pets in the City of Moscow.

Materials and Methods

The study was conducted at the Veterinary Laboratory “Pasteur”. Animal fecal samples were delivered to the laboratory in special plastic containers or glassware.

Feces obtained from various animal species were studied using a variety of methods including flotation with sodium chloride and ammonium nitrate solutions. Some flotation and sedimentation techniques are used only in Russia. For instance, Kotelnikov-Varenichev’s and Kotelnikov-Chrenov’s methods are centrifugal flotation techniques, which have high sensitivity for many helminth species ( Kotelnikov, 1974 ; Kotelnikov & Chrenov 1980 ), ether and formalin precipitation, and direct smear microscopy (MUC 4.2.3145-13). Sometimes all the research methods were used simultaneously based on the particular features of the approach used to identify various species of protozoa, in accordance with the nature of the fecal sample under test.

Microscopy was performed at 10x and 40x magnification using a Lomo microscope (Joint stock company Lomo, Russia). The species of helminth eggs and protozoan cysts were determined based on their morphological features and size.

After examination, the material samples were disinfected by autoclaving.

Ethical Approval

The animal study was in compliance with all the 209 relevant national regulations and institutional policies relating to the care and use of animals in Russia.

The pet animal study was conducted for 6 years from 2012 to 2017. During this period, we studied fecal samples from 1752 dogs ( Canis lupus ) , 1261 cats ( Felis catus ), 323 ferrets ( Mustela putorius ), 217 chinchillas ( Chinchilla lanigera ), 132 guinea pigs ( Cavia porallus ), 165 rabbits ( Oryctolagus cuniculus ), 65 tamarins ( Saguinus oedipus ), 50 marmoset ( Callithrix jacchus ), 10 galago ( Galago senegalensis ), 10 chimpanzee ( Pan troglodytes ), 10 turtles ( Testudo horsfieldii ), 11 geckos ( Gekkonidae ), 21 hedgehogs ( Erinaseus europaeus ).

In the dogs, we discovered eight different helminths and seven different protozoa, i.e. Toxocara canis , Toxascaris leonina , Trichuris vulpis , Ancylostomatidae , Ancylostoma сaninum , Spirocerca lupi , Taenia sp., Capillariа sp.; Sarcocystis sp., Cystoisospora canis , Cystoisospora sp., Giardia sp., Neospora caninum and/or Hammondia heydorni , Thrichomonadidae, and Cryptosporidium sp. ( Table 1 ).

Indices of infection prevalence (IP) for parasitic diseases in pet dogs.

T.canis eggs were found in 43 samples (2.4 % infection prevalence (IP) is the percentage of a infected animals in the total population) ( Fig. 1 )., T.leonina eggs in 8 samples (0.4 % IP), Strongyloides sp. larvae in 16 cases (0.9 % IP) ( Fig. 2 ), Trichuris vulpis in 5 samples (0.28 % IP), S.lupi in 1 sample (0.05 % IP), Taenia sp. in 4 cases (0.2 % IP), Ancylostomatidae in 10 cases (0.5 % IP), A.сaninum in one case (0.05 % IP), and Capillariа sp. in 3 cases (0.17 % IP). Intestinal protozoa including Giardia sp. were identified in 172 cases (9.8 % IP), Sarcocystis sp. in 110 cases (6.2 % IP), Cystoisospora sp. in 36 cases (2 % IP), C.canis in 17 cases (0.9 % IP), Cryptosporidium sp. in 18 cases (1 % IP), N.caninum or/and H.heydorni in 6 cases (0.34 % IP), and Thrichomonadidae in 4 cases (0.2 % IP).

Eggs of Toxocara canis in dog.

Larva of Strongyloides sp. in dog, stage L1.

In the case of mixed infections are most frequently caused by a combination of two species of protozoa, less frequently by a combination of one type of helminths and one type of protozoa. A total of 59 cases of mixed infections in dogs were detected ( Table 4 ).

Combination mixed infections in dogs and cats.

It was determined that young animals (younger than one year of age) had the highest level of intestinal parasite infection in dogs. Thus, the prevalence of toxocarosis in puppies, based on the total number of positive samples (43 samples were positive, with 31 of them obtained from puppies) was 72 %. The incidence was 62.2 % for Giardia sp., 76.4 % for C. canis , 69.4 % for Cystoisospora sp., and 55.5 % for Cryptosporidium sp. The frequency of detection for Sarcocystis sp. was 36.3 %.

In the pet cats, we identified the following 6 species of helminths: T. cati , T. leonina , Capillaria sp., Dipylidium caninum , Ancylostomatidae , Taenia sр. and 6 species of intestinal protozoa: Giardia sp., Cystoisospora rivоlta , Cystoisospora felis , Sarcocystis sp., Thrichomonadidae, and Cryptosporidium sp.( Table 2 )

Indices infeсtion of prevalence (IP) for parasitic diseases in pet cats.

The most common helminthiasis in cats was toxocarosis, with T. cati eggs found in 73 samples (5.7 % infection prevalence (IP )), proglottids of D.caninum in 16 samples (1.2 % IP), Taenia sp. eggs in 10 samples (0.7 % IP), Capillaria sp. in 7 samples (0.5 % IP), T.leonina in 1 (0.07 % IP), and Ancylostomatidae in 2 samples (0.15 % IP).

The most common feline protozoan disease was Giardia sp., which was found in 59 samples (4.6 % IP) ( fig 3 ). C. felis was found in 32 samples (2.5 % IP), C. rivоlta in 11 samples (0.8 % IP), Sarcocystis sp. in 9 samples (0.7 % IP), Cryptosporidium sp. in 15 samples (1.1 % IP), and Thrichomonadidae in 30 samples (2.3 % IP).

Trophozoites and cysts of Giardia sp. in cat.

There were 26 cases of mixed infections in the cats, most of them were caused by two species of parasites, with only one case involving a combination of 4 species of parasites ( Table 4 ).

In the ferrets, we found a single type of helminths belonging to the genus Capillaria (0.3 % infection prevalence (IP) and 3 species of protozoa, i.e. Cryptosporidium sp., Giardia sp., and Eimeria sp. ( Table 3 ). The most frequently seen parasites were Cryptosporidium sp., found in 21 cases (6.5 % IP) ( Fig. 4 ), and Eimeria sp., found in 17 cases (5.2 % IP) ( Fig.5 ), with Giardia sp. cysts represented to a lesser extent. Only one case involving one type of helminths, i.e. H.nana , was identified in the chinchillas along with 4 species of protozoa: Giardia sp., Blastocystis sp., Entamoeba sp., and Trichomonas sp. The most common intestinal protozoa in the chinchillas were Giardia sp. with 193 cases and an IP value of 47.7 %. There were 4 cases of Trichomonas sp. (1.8 % IP) and isolated instances of other species (0.4 % IP). The only intestinal parasites found in the guinea pigs were protozoa represented by three species, i.e. Entamoeba sp., Trichomonas sp., and Giardia sp., with the most frequently seen protozoa being Trichomonas sp. with 12 cases (9 % IP) and cysts of Giardia sp. and amoebas (3.7 % IP). One species of helminths, i.e. Passalurus ambiguus with an IP of 2.4 %, 2 species of Eimeria sp. with 23 cases (13.9 % IP), and Giardia sp. with 3 cases (1.8 % IP), which were encountered much less frequently, were identified in the rabbits. The primates (data for all types) were shown to be 15.8 % infected with Acanthocephala. The stomach nematodes of the Streptopharagus sp. are seen much less frequently: 2.8 %. Intestinal protozoa were represented by 4 species: Giardia sp. (3.5 % IP), Blastocystis sp. (2.1 % IP), Entamoeba sp. (3.5 % IP), and Trichomonas sp. (2.8 % IP). One species of helminths of the genera Oxyurida was identified in four cases in the reptiles along with 4 species of protozoa: Blastocystis sp. (18.1 % IP), Entamoeba sp. (9.5 % IP), Trichomonas sp. (33.3 % IP) and Isospora sp. (14.5 % IP). Only two species of helminths were identified in the hedgehogs, i.e. Capillaria sp. in 7 cases (33.4 % IP) and Crenosoma striatum in 2 cases (9.5 % IP) ( Fig. 6 ).

Indices of prevalence of infeсtion (IP) by intestinal parasites in other house-kept animals.

Oocysts of Cryptosporidium sp. in ferret.

Oocysts of Eimeria sp. in ferret.

Larva of Crenosoma striatum in hedgehog, stage L1.

In certain isolated cases, feces from lemurs ( Lemur catta ) containing helminths of the Enterobius sp. (data not listed in the tables) were submitted for examination along with feces from meerkats ( Suricata suricatta ), raccoons ( Procyon lotor ), degu ( Octodon degus ), and echidna ( Tachyglossidae ), in which no intestinal parasites were identified. Protozoa from the Eimeria sp., eggs of Catenotaeniidae and eggs of the genera Oxyurida were found in squirrels ( Sciurus vulgaris ).

The study has shown that Toxocara spp. are the most widespread parasites in dogs and cats. Different researchers give different IP values, which may be dependent on the category of the animals under study (examination of all animals in veterinary clinics, animals in shelters, stray animals) (Kurnosova & Odoyevskaya, 2017). It is quite common for Toxocara spp. infections to rank first in terms of prevalence in dogs and cats in an urban environment. In addition, cats have a higher level of infection with Toxocara sp. than dogs. Currently the role of cats in the epidemiology of human toxocariasis remains unclear and it may be true that the latter are a source of infection for humans to a greater extent than previously thought ( Fogt-Wyrwaset et al., 2007 ; Espinoza et al., 2010 ; Panova, 2011 ). The results of our study are in agreement with other authors ( Prozorov, 1999 ; Zubareva, 2001 ; Mikhina, 2008 ; Kurnosova, 2009 ; Panova, 2011 ; Kurnosova & Odoyevskaya, 2017 ;). In this situation, the urban environment favors the spread of toxocarosis infection, in which the main factor of accumulation and spread is the ground of the pet walking areas and the permanent habitat of stray dogs and cats. These conditions ensure constant circulation of this species of helminths, which is fraught with constant re-infection and infection of additional animals. The rest of the helminth species are found less frequently, which is due to the conditions of indirect transmission of a particular species involving intermediate and/or additional hosts. In such a case, cats and dogs get infected when they go away from the city for the summer. In recent years, larvae of Strongyloides sp. have been found more frequently than before (Umur & Meral, 2017; Paradies & Iarussi, 2017). This nematode species is found not only in dogs going away from the city or country, but also in animals residing within city limits at all times, mostly in puppies kept in breeding kennels. The Laboratory received material from animals suffering from long-term infection. The course of this type of infection is severe in young animals and disrupts the normal functioning of the gastrointestinal tract, which is manifested in a loss of appetite, copious mucus-laden stools, or intractable diarrheas ( Umur &Meral, 2017 ).

Giardiosis ranks first in terms of prevalence among the protozoan diseases of dogs, cats, and chinchillas, with puppies younger than 1 year of age affected most frequently in the case of dogs (Upjonh & Cobb, 2010; Kurnosova, 2014 ). At the same time, in cats and chinchillas this species of protozoa is found with equal frequency in young and adult animals. Such prevalence of Giardia spp. is related to their environmental resistance ( Adam, 2001 ). In dogs, Giardia spp. are found more frequently than in cats, which may be due to the animal’s daily walking routines and their infection from the environment, as well as to the habit of some dogs to eat the feces of other animals. An animal with weak immunity may become a long-term host to these protozoa. As of now, the role of pet animals in the transmission of this infection to humans within city limits has not been established ( Feng & Xiao, 2011 ; Thompson & Monis, 2004 ; Thompson & Palmer, 2008 ; Upjonh & Cobb, 2010). Sarcocystidae are found in dogs more frequently than in cats, which is due to the fact that cats are more often fed dry kibble. In general, however, the prevalence of Sarcocystidae infection has declined in recent years and they now rank second after giardiosis. Infections by flagellated protozoa from the Thrichomonadidae family have recently started cropping up in cats of various ages. These protozoa are sufficiently pathogenic and difficult to treat. The species of trichomonads circulating among cats within city limits have not been established. The animals suffered from chronic diarrhea whenever trichomonads were found. In dogs and cats, Cryptosporidium sp. and Cystoisospora spp. are less prevalent and are found mostly in young animals( Hamnes & Gjerde, 2007 ; Garanayak & Gupta, 2017). It should be noted that protozoa from the family Sarcocystidae have not been detected in feline fecal samples during the entire study period, which is compatible with the widely expressed view regarding quick and often asymptomatic excretion of cysts into the environment that goes unnoticed by the owners (Greene, 2011).

In general, and as already shown by previously conducted studies, intestinal protozoa in dogs and cats are found more frequently than helminthosis ( Kurnosova, 2009 ).

Of particular interest is research relating to the group of exotic animals, which are not kept in urban apartments as much as other species. Nevertheless, many species of these animals are gaining in popularity. Monkeys, squirrels, and some species of rodents are normally kept in mini-zoos and large shopping malls in Moscow. They are much less frequently kept by individuals. Owners are increasingly starting to keep rabbits and, less frequently, reptiles and hedgehogs in urban apartments. The research has shown that exotic animals are infected mostly by helminths and protozoa found only in these animal species, although some need to be highlighted their importance in Public Health (Hallinger & Taubert, 2018). The primates infected with Acanthocephala. This species of helminths is widespread in animals, which is due mostly to the fact that the animals are fed insects acting as intermediate hosts to these helminths. While the protozoa from the Entamoeba sp. that are most frequently found in monkeys, reptiles, and guinea pigs are represented mostly by Endolimax nana and Iodamoeba sp., it has not been always possible to determine the species of some amoeba cysts. No dysenteric amoeba ( Entamoeba histolytica ) cysts capable of transmission to humans have been identified over the entire study period ( Regan & Yon, 2014 ; Levecke & Dreesen, 2010 ; Verweij & Brienen, 2003 ). However, in addition to the species-specific parasites, helminths and protozoa of a zoonotic nature (eggs of Hymenolepsis nana , blastocysts) have also been found in this category of animals under study (Cian & Safadi, 2017). Certainly, when an exotic animal is bought, the animal should be screened for parasitic diseases. In addition, animals leaving city or country limits or those participating in hunting, actively catching and eating insects, rodents, and amphibians may get infected by zoonotic diseases (Hallinger & Taubert, 2018). The same danger should be kept in mind when brining the younglings of wild animals from the forest.

Thanks to the veterinary and medical awareness-building work by the mass-media and the Internet, pet owners have become sufficiently knowledgeable about many issues relating to animal diseases and various preventative activities. Nevertheless, studies have shown that under urban conditions parasitic diseases are relevant both to veterinary science and to medicine ( Stepanov, 2014 ; Shcheveleva et al., 2016 ).

Thus, veterinarians and pet owners should take the species of their animal, it’s origin, living and feeding conditions into consideration in order to take preventative measures aimed at timely identification of parasitic infections and their correct treatment under veterinary supervision, which will ensure the animal’s health and protect the environment from the spread of infections. Studying a wide range of pet animals under urban conditions has great value not only from a practical standpoint, but also in terms of determining the epizootic situation for all the major parasitic diseases in general and clarifying the human risks associated with the zoonotic nature of the parasitic infections.

The study has shown the widespread circulation of intestinal parasitic infections of house-kept animals in the city of Moscow. The high concentration of animals within confined areas promotes constant circulation of numerous helminths and protozoa among pet animals. Тoxocara spp. and Giardia sp.were found more frequently in dogs and cats together with other parasitic agents are a pressing issue both for veterinary science and for medicine. In summary, helminths and protozoa capable of transmission to humans have been identified among a wide range of pet animal infections in an urban environment.

Conflict of Interest

Authors state no conflict of interest.

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