Otter (Lutrinae) Care Manual - Association of Zoos and Aquariums

Otter (Lutrinae) Care Manual 

Created by the 

AZA Small Carnivore Taxon Advisory Group 

in Association with the 

AZA Animal Welfare Committee 

1


Published by the Association of Zoos and Aquariums 

Association of Zoos and Aquariums 2 


Formal Citation: 

AZA Small Carnivore TAG 2009. Otter (Lutrinae) Care Manual. Association of Zoos and Aquariums, 

Silver Spring, MD. 

Original Completion Date: 

Original version 2002, 4 th Revision October 2009 

Authors and Significant contributors: 

Jan Reed Smith (Columbus Zoo and Aquarium), Celeste (Dusty) Lombardi (Columbus Zoo and 

Aquarium), Kim Lengel (Philadelphia Zoo), Mike Maslanka (Smithsonian National Zoological Park), Barb 

Henry (Cincinnati Zoo), Gwen Myers, D.V.M. (Columbus Zoo and Aquarium), Contributors: Jessica Foti 

(North Carolina Zoological Park), Juan Sabalones (Maryland Zoo), Sheila Sykes-Gatz (Dortmund Zoo). 

AZA Staff Editors: 

Lacey Byrnes, B.S. Animal Care Manual Intern 

Candice Dorsey, Ph.D., Director, Animal Conservation 

Reviewers: 

AZA Small Carnivore TAG members 

AZA Animal Health Committee 

AZA Nutrition Advisory Group 

Dr. Merav Ben-David, Ph.D. (University of Wyoming) 

Grace Yoxon (International Otter Survival Fund) 

Joseph C.E. Barber Ph.D. (AZA, ACM Consultant) 

Debborah Colbert Ph.D. (AZA, Vice President of Animal 

Conservation) 

Candice Dorsey Ph.D. (AZA, Director of Animal Conservation) 

Paul Boyle Ph.D. (AZA, Senior Vice President of Conservation 

and Education) 

Cover Photo Credits: 

Asian small-clawed otter family: Jennifer Brink, Zoo Atlanta: 

North American river otter group: Doug Kjos, friend of Roosevelt 

Park Zoo; Giant otter and cub: Cali Zoo, Columbia; 

Spotted-necked otter: Jenna Kocourek, Little Rock Zoo 

Disclaimer: This manual presents a compilation of 

©Jason Theuman, Buttonwood Park Zoo 

knowledge provided by recognized animal experts based on the 

current science, practice, and technology of animal 

management. The manual assembles basic requirements, best practices, and animal care 

recommendations to maximize capacity for excellence in animal care and welfare. The manual should be 

considered a work in progress, since practices continue to evolve through advances in scientific 

knowledge. The use of information within this manual should be in accordance with all local, state, and 

federal laws and regulations concerning the care of animals. The recommendations are not exclusive 

management approaches, diets, medical treatments, or procedures, and may require adaptation to the 

specific needs of individual animals and particular circumstances in each institution. Commercial entities 

and media identified are not necessarily endorsed by AZA. The statements presented throughout the 

body of the manual do not represent standards of care unless specifically identified as such in clearly 

marked sidebar boxes.



Table of Contents 

Introduction ...............................................................................................................................................5 

Taxonomic Classification......................................................................................................................5 

Genus, Species, and Status.................................................................................................................5 

General Information...............................................................................................................................5 

Chapter 1. Ambient Environment ........................................................................................................8 

1.1 Temperature and Humidity ............................................................................................................8 

1.2 Light...................................................................................................................................................9 

1.3 Water and Air Quality....................................................................................................................10 

1.4 Sound and Vibration .....................................................................................................................12 

Chapter 2. Habitat Design and Containment ..................................................................................13 

2.1 Space and Complexity..................................................................................................................13 

2.2 Safety and Containment...............................................................................................................18 

Chapter 3. Transport .............................................................................................................................21 

3.1 Preparations...................................................................................................................................21 

3.2 Protocols.........................................................................................................................................22 

Chapter 4. Social Environment...........................................................................................................24 

4.1 Group Structure and Size ............................................................................................................24 

4.2 Influence of Other Species and Conspecifics...........................................................................27 

4.3 Introductions and Reintroductions ..............................................................................................28 

Chapter 5. Nutrition ...............................................................................................................................30 

5.1 Nutritional Requirements..............................................................................................................30 

5.2 Diets ................................................................................................................................................34 

5.3 Nutritional Evaluations..................................................................................................................37 

Chapter 6. Veterinary Care ..................................................................................................................38 

6.1 Veterinary Services.......................................................................................................................38 

6.2 Identification Methods...................................................................................................................38 

6.3 Transfer Examination and Diagnostic Testing Recommendations........................................39 

6.4 Quarantine......................................................................................................................................39 

6.5 Preventive Medicine......................................................................................................................41 

6.6 Capture, Restraint, and Immobilization......................................................................................47 

6.7 Management of Diseases, Disorders, Injuries and/or Isolation..............................................48 

Chapter 7. Reproduction......................................................................................................................53 

7.1 Reproductive Physiology and Behavior.....................................................................................53 

7.2 Artificial Insemination....................................................................................................................64 

7.3 Pregnancy and Parturition ...........................................................................................................65 

7.4 Birthing Facilities ...........................................................................................................................65 

7.5 Assisted Rearing ...........................................................................................................................65 

7.5 Contraception.................................................................................................................................73 

Chapter 8. Behavior Management .....................................................................................................75 

8.1 Animal Training..............................................................................................................................75 

8.2 Environmental Enrichment...........................................................................................................78 

8.3 Staff and Animal Interactions ......................................................................................................81 

8.4 Staff Skills and Training................................................................................................................82 

Chapter 9. Program Animals...............................................................................................................83



9.1 Program Animal Policy .................................................................................................................83 

9.2 Institutional Program Animal Plans ............................................................................................85 

9.3 Program Evaluation ......................................................................................................................86 

Chapter 10. Research............................................................................................................................87 

10.1 Known Methodologies................................................................................................................87 

10.2 Future Research Needs.............................................................................................................88 

Acknowledgements...............................................................................................................................91 

References...............................................................................................................................................92 

Appendix A: Accreditation Standards by Chapter......................................................................100 

Appendix B: Acquisition/Disposition Policy.................................................................................103 

Appendix C: Recommended Quarantine Procedures ................................................................107 

Appendix D: Program Animal Position Statement......................................................................109 

Appendix E: Developing an Institutional Program Animal Policy ..........................................111 

Appendix F: AZA Accreditation Standards for Otters................................................................115 

Appendix G: Giant Otter Enclosure Design ..................................................................................117 

Appendix H: Description of Nutrients ............................................................................................119 

Appendix I: Pup Weights of Ex-situ Population Bred Otters....................................................122 

Appendix J: List of Commonly Trained Behaviors for Otters..................................................124 

Appendix K: Enrichment Items Commonly Provided to Otters...............................................126 

Appendix L: Resources for Enrichment and Training ...............................................................128 

Appendix M: Missouri Fish and Wildlife Otter Stomach Contents .........................................131 

Appendix N: Basic Considerations in the Design and Maintenance of Otter Exhibit Life 

Support Systems..................................................................................................................................132 

Appendix O: Female Otter Reintroduction Plan ..........................................................................153 

Appendix P: Otter Body Condition Matrix.....................................................................................154



Introduction 

Preamble 

Association of Zoos & Aquariums (AZA) accredited institutions are required to comply with all relevant 

local, state, and federal wildlife laws and regulations in addition to all AZA accreditation standards. 

AZA accreditation standards are continuously being raised or added. Staff from AZA-accredited 

institutions are required to know and comply with all AZA accreditation standards, including those most 

recently listed on the AZA website (www.aza.org) which might not be included in this manual. 

Taxonomic Classification 

Table 1: Taxonomic classification for Lutrinae 

Classification Taxonomy Additional information 

Kingdom Animalia 

Phylum Chordata 

Class Mammalia 

Order Carnivora 

Suborder Caniformia 

Family Mustelidae 

Sub Family Lutrinae Some species names are still being debated 

Genus, Species, and Status 

Table 2: Genus, species, and status information for Lutrinae recommended for management by the AZA Otter SSP 

Genus Species Common 

Name 

USA Status IUCN Status AZA Status 

Aonyx (Amblonyx) Cinereus Asian small- Not listed Vulnerable 

SSP 

(cinerea)* clawed otter 

Lutra (Hydrictis) 

maculicollis Spotted-necked Not listed Least Concern SSP 

Pteronura 

otter 

brasiliensis Giant otter Endangered Endangered SSP 

Lontra (Lutra) canadensis North American Not listed Least Concern PMP 

river otter 

*Some recent resources are still using Aonyx cinerea (Wilson & Reeder 2005) however, the IUCN/SSC Otter Specialist Group and 

ITIS have switched to Aonyx cinereus based on input from Koepfli & Wayne (1998, 2003) and advice that the appropriate Latin 

gender declination is cinereus and not cinerea; we are using the OSG as our citing source (www.otterspecialistgroup.org/). ISIS 

uses A. cinereus. 

General Information 

Introduction: The AZA Small Carnivore Taxon Advisory Group (TAG) has designated four semi-aquatic 

otter species for management under the AZA Otter Species Survival Plan® (SSP). These are the Asian 

small-clawed otter (Aonyx/Amblonyx cinereus), giant otter (Pteronura brasiliensis), North American 

(Nearctic) river otter (Lontra canadensis), and the spotted-necked otter (Lutra maculicollis). The African 

(Cape) clawless otter (Aonyx capensis) is currently represented in AZA institutions but is recommended 

for phase-out. Because specimens are still maintained by member institutions, care information is 

included. These otter species exhibit varying needs and degrees of sociality. Many of their ex-situ 

population care requirements are similar but there are significant variations in some of their husbandry 

needs. For more detailed information please refer to the following husbandry manuals: 

• Asian Small-clawed Husbandry Manual (Lombardi et al. 1998) 

• International Giant Otter Studbook Husbandry and Management Information and Guidelines 

2005, 2 nd Edition (Sykes-Gatz 2005) 

• North American River Otter Notebook, 3rd Edition (Reed-Smith 2008) and Summary of 

Husbandry Guidelines for North American (aka Nearctic) River Otters (Lontra canadensis) in 

Captivity (Owens et al. 2009): www.otterspecialistgroup.org/Library/TaskForces/OCT.html. 

• Summary of Husbandry Guidelines for African Spotted-necked Otters in Captivity (Benza et al. 

2009): www.otterspecialistgroup.org/Library/TaskForces/OCT.



Natural History/Description: 

Asian small-clawed otter (Aonyx cinereus): This is one of five otter species found in Asia. It is one of the 

smallest of the world’s otters, rarely weighing more than 5kg. A gregarious species, it is often seen in 

large groups of up to 15 animals, and ex-situ population studies suggest that these groups are composed 

of an alpha breeding pair and their offspring from successive litters. Asian small-clawed otters have 

unusual hand-like front paws with increased tactile sensitivity and reduced webbing, which they use to 

forage for their prey of crustaceans, mollusks, and small fish. 

Asian small-clawed otters are found from Palawan (Philippines) through Indonesia, Southeast Asia, 

southern China, and westwards throughout the Himalayan foothills of Bangladesh, Bhutan, and Nepal. A 

disjunctive population occurs in southern India (Foster-Turley et al. 1990). Listed on CITES as Appendix II 

(www.cites.org) and as Vulnerable by IUCN/SSC the population is considered to be decreasing. 

African (Cape) clawless otter (Aonyx capensis): The African clawless otter is one of four species of otters 

found in Africa. It is the third largest species of otter. Only the giant otter and sea otter are larger. Adults 

range in size from 1.15-1.5m (3.8-5ft), and weigh from 16-20kg (35.3-44.1lbs) (Foster-Turley et al. 1990). 

The African clawless otter has been reported as living in family groups including the male, female, and 

pups (Rowe-Rowe 1978), family groups consisting of the female and pups, or singly (Chanin 1985). The 

prevalent social grouping may vary with the habitat, which also likely influences the size and degree of 

overlap of home ranges. African clawless otters use their sensitive, non-webbed fingers to forage food, 

which consists primarily of fresh-water crabs, crayfish, and some fish. In some areas, this species is 

reported to occasionally raid near-by farms for young maize and cabbages (J.Reed-Smith, personal 

communication). 

This species is distributed from Ethiopia in the east to Senegal in the west and south to South Africa, 

with a distributional gap in the rain forest area of the Congo basin, where the Congo clawless otter is 

found (Rowe-Rowe 1991). The African clawless otter is found in both fresh water streams and rivers, and 

along marine coastlines in South Africa. 

Due to the infrequency of the holding and exhibition of the African clawless otter in zoos and 

aquariums, many of the standards set for this species are extrapolated from those set by the N.A. river 

otter and Asian small-clawed otter husbandry manuals. Listed on CITES as Appendix II (www.cites.org). 

Giant otter (Pteronura brasiliensis): This single species in the genus Pteronura is one of four species of 

otter found in South America. The giant otter’s large size (1.5-2m, 4.9-6.6ft), weight (25-32kg, 55.1- 

70.5lbs) (Duplaix 1980), highly social nature (multi-generational family groups), and critically endangered 

status make this species attractive to many facilities. However, their specific housing requirements and 

sensitivity to disturbances make them one of the more difficult otter species to hold successfully in zoos 

and aquariums. The diet of the giant otter is comprised almost completely of fish. 

Although originally found in Colombia, Venezuela, Guyana, French Guiana, eastern Ecuador, Peru, 

Brazil, Bolivia, Uruguay, Paraguay, Suriname and northeastern Argentina, only remnant populations of 

giant otter are currently found throughout its former range. It is mainly found in slow moving rivers and 

creeks within forests, lakes, ox-bow lakes, swamps, and marshes in the tropical lowland areas of South 

America. With an estimated total population of only 1,000-5,000 individuals, the giant otter is considered 

highly vulnerable to extinction. It is classified as Endangered by the World Conservation Union (IUCN), as 

Endangered by the US Fish and Wildlife Service, and is listed on Appendix I of the Convention on 

International Trade in Endangered Species (CITES). Historically hunted for pelts, the species is now 

threatened by increased human colonization of tropical lowland rainforests. Other threats include habitat 

destruction and degradation, over-fishing, illegal hunting, mining, and water and land pollution. 

North American river otter (Lontra canadensis): The North American river otter is one of the four new 

world river otter species. There are at least seven subspecies of L. canadensis. Adults range in size from 

1-1.53m (3.3-5ft) and weigh from 4.5->16kg (9.9-35.2lbs) (Ben-David et al. 2001a,b; Reed-Smith 2001). 

Although frequently solitary, except for female with pups, the North American river otter shows a great 

deal of social plasticity (particularly males), often forming groups of 8-15 or more animals in environments 

offering abundant resources (Blundell et al. 2002a,b). All male groups of up to 15 individuals have been 

maintained successfully in zoos and aquariums (Ben-David et al. 2000). In the wild, males do not 

participate in pup rearing; in zoos and aquariums males can be reintroduced to the family group once the 

pups are swimming well and in general interact and play with the pups. Both sexes occupy linear shaped 

home ranges due to their affinity for the land/water interface. Activity centers (e.g., latrines), located within



home ranges, are important for both sexes. During a latrine activity study in Pennsylvania, Stevens & 

Serfass (2008) documented that visiting otters spent 72.7% of their time there smelling and investigating, 

10.9% marking, 10.6% traveling, 4.6% rolling and rubbing, and

1.1 Temperature and Humidity 

Air Temperature: All otter species should be provided with 

shelter from the sun and inclement weather. Indoor exhibits 

should offer an ambient temperature gradient within the exhibit 

providing otters the opportunity to select for their comfort. 



Chapter 1. Ambient Environment 

AZA Accreditation Standard 

(1.5.7) The animal collection must be 

protected from weather detrimental to 

their health. 

A. cinereus: The ideal air temperature is between 22.2-24.4°C 

(72-76˚F). If Asian small-clawed otters have access to radiant heat, or a heated indoor facility, they can 

handle temperatures down to 10-15°C (50°F). The recommended water temperature is between 18.3- 

29.4°C (65-85˚F). It is recommended that warm water (29.4°C/85˚F - Lombardi 2004) be provided for 

swimming, since these tropical animals will spend more time in the water if it is warm (Petrini 1998), and 

this may have beneficial health effects. 

A. capensis and L. canadensis: These species can tolerate a wide temperature range as long as they are 

offered protection from the sun and inclement weather in outdoor exhibits. Indoor exhibits should offer a 

thermal gradient allowing animals the selection of a comfortable temperature (10-24°C or 50-75˚F) (Reed- 

Smith 2004a). A temperature below 21-24°C (70-75˚F) is recommended for indoor holding/night facilities 

(Wallach & Boever 1983). Animals should always be provided with shelter from the sun in outdoor 

exhibits. 

L. maculicollis: This species has been housed successfully at floor temperatures ranging from 14.4- 

25.5°C (58-78°F) (Schollhamer 1987). Their temperature tolerance is likely to be similar to that of A. 

cinereus and A. capensis; however, at this time there is insufficient information and experience to make 

informed recommendations. Animals should be monitored for signs of overheating and hypothermia at 

temperatures above 25.5°C (78°F) and below 14.4°C (58°F), respectively. 

P. brasiliensis: In temperate climates, Wünnemann (1995a) recommends a minimum of 18°C (64.4°F) air 

temperature for dens and indoor enclosures. The suggested temperature range is 18-20°C (64.4-68°F) 

(Hagenbeck & Wünnemann 1992). For specific temperature recommendations for young pups, see 

Chapter 7. Indoor enclosures should be equipped with fans, cooling, and/or ventilation systems to prevent 

over-heating and provide fresh air exchange in all climates; in temperate climates a heating system is 

required (Sykes-Gatz 2005). 

This species should be provided with the choice to use an outdoor enclosure year-around, even in 

temperate climates, as they are quite adaptable to colder outdoor temperatures (young pups are an 

exception to this, see Chapter 7), as long as they have access to heated indoor enclosures in addition to 

their dens (Wünnemann 1995a). Adults will not carry out their normal daily terrestrial activities in air 

temperatures at approximately 10°C (50°F) or below (regardless of whether the outdoor pool water is 

heated), but will spend limited time in these temperatures, and seem to avoid temperatures that are too 

cold for them. Giant otters should have access to a heated indoor enclosure at all times when seasonal 

daytime air temperatures regularly fall below 15°C (59°F) (Sykes-Gatz 2005; V.Gatz, personal 

communication). The following recommendations are provided for giant otters: 

• Exposure to air temperatures at or below ~7°C (20°F) should be restricted, and otters should be 

carefully monitored if given access to temperatures near this range. 

• Newly imported animals from tropical climates, juveniles, and sub-adults should be acclimated 

slowly over a period of 6-12 months to these colder temperatures. 

• Shelter from the wind, rain, heat, cold, and constant direct sun should be provided in all climates 

(Sykes-Gatz 2005). 

Water Temperature: More detailed research is required into optimal water temperature levels for the 

tropical otter species; however, at this time the AZA Otter SSP recommends the following temperature 

guidelines: 

A. cinereus: The water temperature for A. cinereus should be maintained between 18.3-29.4°C (65-85°F), 

preferably at the warmer end of this scale (Petrini 1998).



A. capensis, L. canadensis: The water temperature for A. capensis and L. canadensis does not appear to 

be critical. 

L. maculicollis: Water temperature in successful L. maculicollis exhibits has ranged from 8.9-15.6°C (48- 

60°F). Temperatures in the 15.6-21.1°C (60-70°F) range may encourage this species to spend more time 

in the water, however, this is has not been objectively demonstrated at this time. 

P. brasiliensis: Further study into optimal pool temperatures and water temperature exposure 

recommendations for P. brasiliensis is required. Sykes-Gatz (2005) recommends this species should not 

be allowed to swim in unheated water when air temperatures are below 5°C (41°F). As a precaution, 

outdoor pools should be emptied when temperatures approach this range. Sufficient indoor swim areas 

are needed when seasonal daytime air temperature regularly falls below 15°C (59°F), regardless of 

whether outdoor water is heated. This is particularly true for family groups rearing pups that may be held 

indoors for 4-5 months during cold temperatures. Heating of indoor housing pools is not necessary if the 

ambient air temperature is maintained at recommended levels. See Appendix G for information on pool 

design recommendations for giant otters. 

Humidity: Since otters always should have water features available to them, humidity does not seem to 

be a factor in their environment unless it is excessive. Excessive humidity and an inability to adequately 

dry off create problems for all otter species, and these conditions should be avoided. The relative 

humidity of indoor exhibits should range between 30-70%. Nest boxes and den sites should be provided 

with good ventilation and placed in locations that are not chronically humid. The AZA Otter SSP 

recommends the provision of sufficient dry land (see Chapter 2, section 2.1), natural substrates, and 

bedding material (see Chapter 2, section 2.1) to aid the otters in 

proper coat maintenance, and allow for adequate drying of their 

pelts and feet. 

AZA institutions with exhibits which rely on climate control 

must have critical life-support systems for the animal collection 

and emergency backup systems available, while all mechanical 

equipment should be included in a documented preventative 

maintenance program. Special equipment should be maintained 

under a maintenance agreement or records should indicate that 

staff members are trained to conduct specified maintenance 

(10.2.1). Records should include daily activities required for 

maintenance, such as back-washing of filtration systems, ozone 

checks, pH, chlorine, and coliform levels as well as dates of 

periodic maintenance activities. 


(10.2.1) Critical life-support systems for 

the animal collection, including but not 

limited to plumbing, heating, cooling, 

aeration, and filtration, must be equipped 

with a warning mechanism, and 

emergency backup systems must be 

available. All mechanical equipment 

should be under a preventative 

maintenance program as evidenced 

through a record-keeping system. Special 

equipment should be maintained under a 

maintenance agreement, or a training 

record should show that staff members 

are trained for specified maintenance of 

special equipment. 

1.2 Light 

Careful consideration should be given to the spectral, intensity, and duration of light needs for all 

otters in the care of AZA-accredited zoos and aquariums. 

If otters are held in holding areas for any length of time, some natural light is recommended. 

Fluorescent, metal halide and mercury vapor, as well as natural light have all been used in exhibit areas. 

The AZA Small Carnivore TAG is unaware of any hard data on the impact of light intensity on otter health 

or reproduction; this should be investigated in the future. However, it is strongly suspected that otters held 

indoors should be provided a seasonally appropriate light cycle to promote breeding and general animal 

health (Bateman et al. 2009). There are no available data on possible deleterious effects of less than full 

spectrum light on a long-term basis. 

A. cinereus: If housed indoors, these species should be kept on a 12-hour light cycle (Wilson, Tropea & 

Calle, unpublished data). 

L. canadensis, A. capensis, L. maculicollis: The light cycle for indoor exhibits/holding should be set to 

mimic the natural photoperiod for the species range of origin in Equatorial Africa, for A. capensis and L. 

maculicollis, or the local photoperiod for N.A institutions housing L. canadensis (Reed-Smith 2001, 

Bateman et al. 2009). 

P. brasiliensis: All indoor enclosure areas, except for the nest boxes, should be kept on a 12-hour light 

cycle to mimic the natural habitat conditions of giant otters. If possible, full-spectrum lighting should be



provided. Giant otters are diurnal, and only the nest boxes should remain dark. If necessary, infrared 

lighting may be used when video cameras do not have infrared capabilities. 

1.3 Water and Air Quality 

AZA-accredited institutions must have a regular program of 

monitoring water quality for collections of aquatic animals and a 

written record must document long-term water quality results and 

chemical additions (1.5.9). Monitoring selected water quality 

parameters provides confirmation of the correct operation of 

filtration and disinfection of the water supply available for the 

collection. Additionally, high quality water enhances animal health 

programs instituted for aquatic collections. 


(1.5.9) The institution must have a regular 

program of monitoring water quality for 

collections of fish, pinnipeds, cetaceans, 

and other aquatic animals. A written 

record must be maintained to document 

long-term water quality results and 

chemical additions. 

Air Changes: The number of air changes per hour of non-re-circulated air needed to control odors and 

maintain a healthy condition for the animals and public will vary according to the number of animals in the 

enclosure and the size/volume of the enclosure. The initial design should be for the maximum number of 

animals that could be housed in that particular enclosure. Standardized rates of change for various 

human-occupied enclosures suggest that pet shops require a rate of air exchange of non-recirculated air 

equal to 1 cubic foot of air/minute/ft 2 of floor space, in order to keep odors down to a level acceptable by 

the public (Anon. 1981). Pupping dens may well need higher rates of air exchange in order to maintain air 

quality and/or low humidity. It should be noted, however, that no work has been done specifically 

targeting air change rates for otter exhibits or dens. 

As a general rule, indoor exhibits should have a negative air pressure of 5-8 air changes per hour of 

non-recirculated air. Glass barriers and separate ventilation systems between indoor exhibits and visitor 

areas (Moore 1997) will help reduce the potential of disease transmission from the public as well as 

complaints due to odor. 

Water Quality: See Appendix N for information on designing a life support system for otter exhibits, 

glossary of terms, and additional details on water treatment provided by J. Sabalones (2009) life-support 

systems advisor to the IUCN Otter Specialist Group’s Otters in Captivity Task Force. 

Otters are semi-aquatic mammals, using bodies of water for foraging, transportation corridors, mating 

(typically), cleaning, and “play-type” behavior. It is recommended to monitor nutrients and perform pool 

water changes as needed. There are no standards yet established for pools provided to semi-aquatic 

otters, however, it is suggested that coliform levels be maintained at 400 per ml water or lower, which is 

the standard set for seal rehabilitation pools. A level of 100 per ml is considered safe for humans. All 

chemical additives should be monitored daily and recorded. 

It is recommended that filtration be used in closed otter pools. Sand filters, pool pumps, charcoal 

filters, and ozone pressure sand filters have all been used effectively. Ultraviolet sterilization has proven 

helpful in inhibiting algae build-up, particularly when combined with regular cleaning of pool-sides. Drain 

outlets, filters, and skimmers should be covered or designed to prevent furnishings from blocking them, or 

otters from becoming stuck in them. Daily use of long-handled skimmers will help remove floating debris 

and keep skimmers open and flowing. Natural flow-through systems also work well in otter exhibits, as 

long as the water is determined to be clean and free of heavy pollutants. In general, otters should be kept 

in fresh water systems; however, Ben-David et al. (2000, 2001a,b) successfully kept a group of 15 males 

in sea water that was changed daily. In this case, fresh water was provided in tubs for the animals to 

bathe in. Regardless of water treatment method used, an additional source of fresh, potable drinking 

water should be available at all times.



Accreditation Standards and Related Policies 

The provision of fresh potable water is a requirement of USDA Animal Welfare Regulations (AWR 2005) 

as stated: “If potable water is not accessible to the animals at all times, it must be provided as often as 

necessary for the health and comfort of the animal. Frequency of watering shall consider age, species, 

condition, size, and type of the animal. All water receptacles shall be kept clean and sanitary” (AWR 

2005). Considering the needs of otters, the AZA Small Carnivore TAG states that otters should be given 

fresh water daily if their pools are not filtered or dumped and filled on a daily basis. AZA Accreditation 

Standards require that institutions abide by relevant federal laws and regulations: “The institution must 

comply with all relevant local, state, and federal wildlife laws and regulations. It is understood that, in 

some cases, AZA accreditation standards are more stringent than existing laws and regulations. In these 

cases the AZA standard must be met” (AZA 2008). 

Water quality should be maintained at a level sufficient to control bacterial counts and organic load, 

and to allow clear underwater visibility of animals for health inspections. Clarity and color maintained to 

provide a perceived color of clear and/or blue water is preferred by most facilities. This water clarity is an 

aesthetic requirement only, as long as the water quality is maintained, and the presence of floating algae 

or other material is not harmful to the otters. Otters can be messy eaters and will track a lot of particulate 

debris into their pools. All food remains should be removed from pools daily to prevent consumption of 

spoiled items. The filtration system needs an effective means of skimming (from top to bottom) particulate 

matter. Turn over rate using rapid sand filtration is suggested to be once/hour; meaning that the total 

exhibit water volume should be turned over at least one time per hour when using rapid sand filtration. 

When using rapid sand filtration couple with Ozone; the turnover rate can be extended to once every 1.5 

hours. 

Chemical treatment such as ozone applied to foam fractionation is recommended for marine 

systems. Ozone applied through a contact chamber in conjunction with a low dosage of chlorine is an 

effective treatment for freshwater systems. A large surface area biological filter bed should be 

incorporated if possible. This will allow a natural nutrient removal system to establish itself, which will 

provide system stability. It also will help reduce organic loading as well as reduce colonization of 

undesirable bacteria species. The tank effluent should be pre-filtered before it is sent to the sand filters 

and foam fractionators. 

At least bi-weekly water quality tests are recommended for bacterial counts and daily tests of 

chemical additive levels. Records should be maintained and available for APHIS inspection and reference 

if problems arise. 

Coliform bacteria: Coliform bacterial counts are used to monitor filtration system efficiency and keep track 

of potentially harmful bacteria. Coliform counts should be done at least every other week and more often 

if there are multiple animals using the pool (a policy regarding coliform testing should be set by the 

institution). Often a MPN (Most Probable Number) per 100ml is given as an acceptable limit. However, a 

more accurate measure is the total or fecal coliform count (NOAA 2006). There are no standards yet 

established for fresh-water otter pools. At this time, it is suggested that coliform levels be maintained at or 

lower than levels established for rescued pinnipeds by NOAA. These are: 

• Total coliform counts should not exceed 500 per 100ml water, a MPN of 1000 coliform bacteria 

per 100ml water. 

• Fecal coliform count should not exceed 400 per ml. 

If animal caretakers are routinely exposed to pool water, an institution may establish a higher 

standard of 100 per ml, which is the level considered safe for humans; this should be based on 

institutional policy. 

Chlorine: Many municipalities add chlorine to their water, and readings from tap water of 1ppm or higher 

are possible. While otters generally show no adverse effects from these levels, it is not known what the 

overall impact is to their health and the water repellency of their coats. For this reason, the AZA Otter 

SSP recommends that otters should not be exposed to chlorine levels higher than 0.5ppm for prolonged 

periods, and ideally chlorine should be kept at a non-detectable level. The addition of sodium thiosulfate 

will neutralize any residual chlorine (see below and Appendix M).



Algae control: Algae control is a continuing problem in otter pools, particularly those exposed to 

significant sunlight. There are several techniques that have been used with varying success (see 

Appendix M). 

• Liquid copper sulfate: Liquid copper sulfate can be added directly to the pool water without harm 

to the animals. While this does not get rid of algae, it will inhibit algal growth. 

• UV sterilization: Ultra violet sterilization has proven helpful in inhibiting algae build-up, particularly 

when combined with regular cleaning of pool sides. 

• Chlorine: Chlorine can be used, if necessary, when the otters are not present. In this case, 

sodium thiosulfate can be added and run through the system for an hour before the otters are 

allowed access again. A 5% concentration of sodium thiosulfate added has been successful. To 

obtain the amount needed for a particular system, multiply 0.53 by the volume of the pool in 

gallons; this provides the amount of sodium thiosulfate required in millimeters (C. Harshaw, 

personal communication). 

• Barley straw: Hanging a bag of barley straw in the water stream assists in filtering out algae. This 

should be hung where the otters cannot get to it (e.g., at the top of a waterfall, etc.). Reports of its 

efficacy vary and may be dependent on design and location of pool. 

All of these techniques should be accompanied with routine scrubbing of the pool sides to inhibit 

algae growth, and discussed with life-support professionals. 

Drinking Water: Clean drinking water should be available at all times. Drinking water should be provided 

in bowls small enough that the otters do not swim in them, or via lixits or similar devices. Animals should 

be introduced to the use of lixits (or other drinking fixtures) and monitored by staff until they are certain 

they are proficient in their use. 

Further research is needed into the impact, if any, of pH on otters. The deleterious effects of chlorine 

on otters specifically is anecdotal, e.g. stripping of water repellency; presumed, e.g. potential carcinogenic 

by-products of chlorine break-down; and, unknown, e.g. impact on overall health. While research is 

desirable it is not recommended on these wildlife species and instead caution should be exercised when 

using chlorine in otter pools. 

1.4 Sound and Vibration 

Consideration should be given to controlling sounds and vibrations that can be heard by otters in the 

care of AZA-accredited zoos and aquariums. While there is no evidence that low level background noise 

is disruptive to otters, loud noises can be frightening to them, and high-pitched, long-term noise should be 

avoided. Parturient females should not be subjected to loud or unusual noises; this is particularly true with 

primiparous females and giant otters. Females about to give birth and with very young pups should not be 

subjected to close proximity of the public, loud or unfamiliar voices, construction noise, sudden loud 

noises such as sirens, unfamiliar ambient noises, or vibrations to which they are not already accustomed. 

For the giant otter, both minor and loud sounds that they have been accustomed to before parturition, 

including familiar noises from zoo staff during daily caretaking routines, can also cause significant stress 

after parturition. All precautions should be taken to eliminate these during the last two weeks prior to birth 

(time margin is estimated based on difficulty of predicting parturition dates in these species) and during 

roughly the first month after parturition for primiparous females and for giant otters until pups emerge from 

the den or until roughly 75 to 120 days post parturition when giant otters have demonstrated more noise 

tolerance in the past. 

Otters’ hearing is considered to be good but nothing is known definitively about their hearing acuity or 

frequency ranges heard. Both of these are areas needing further research.

2.1 Space and Complexity 

Careful consideration should be given to exhibit design so 

that all areas meet the physical, social, behavioral and 

psychological needs of the species. Animals should be displayed, 

whenever possible, in exhibits replicating their wild habitat and in 

numbers sufficient to meet their social and behavioral needs 

(1.5.2). 

Important factors to consider when creating successful otter 

exhibits include: exhibit land area size, design and complexity; 



Chapter 2. Habitat Design and Containment 


(1.5.2) Animals should be displayed, 

whenever possible, in exhibits replicating 

their wild habitat and in numbers sufficient 

to meet their social and behavioral needs. 

Display of single specimens should be 

avoided unless biologically correct for the 

species involved. 

pool size--design and complexity (including shoreline length and complexity); substrate materials and 

depths; water quality; climbing surfaces; digging areas; and denning sites (location and construction). 

Land/Water Ratio: Suggested optimal land/water ratios will change as an exhibit size increases or 

decreases. The ratios offered here are for the recommended minimum exhibit size. Smaller exhibits will 

require a higher land area proportion within the ratio. Larger exhibits and P. brasiliensis exhibits 600m ² 

(6,458ft²) in size or more, may have a somewhat lower land proportion and still be successful. . 

L. canadensis, L. maculicollis, and A. capensis: The recommended land/water ratio for L. canadensis, L. 

maculicollis, and A. capensis is 3:1 to 4:1 (3:1 is the absolute minimum land area proportion and 

considered adequate only if the exhibit is large, vertically complex, and offers hard-surface features within 

the pool, such as logs, islands, etc) (Duplaix-Hall 1975; Reed-Smith 2001 2004a). 

A. cinereus: For A. cinereus, the recommended ratio is 5:1 or 6:1 (Duplaix-Hall 1975; Lombardi et al. 

1998). 

P. brasiliensis: P. brasiliensis indoor and outdoor enclosures between 240m² to 600m² (2,583ft² to 

6,458ft²) in size should be provided with an absolute minimum of 60% land area. (Sykes-Gatz 2005; 

Duplaix-Hall 1972 & 1975). As exhibits and any living areas that may include water features decrease in 

size below 240m 2 (2,583ft²), a greater land proportion within the ratio is recommended. E.g. a 150m² 

(1,615ft²) enclosure should provide an absolute minimum of 69% land area and a 75m² (807ft²) enclosure 

needs at least 76.5% land area. Sykes-Gatz (2005) (also see Appendix G) offers a simple formula that 

should be used for guiding land-water calculations. 

Exhibit Complexity – Terrestrial: Otters are land mammals that swim; they are semi-aquatic or 

amphibious, not aquatic animals. Behaviorally healthy otters kept in appropriate enclosure conditions 

spend more of their daytime hours on land than in the water. As 

instinctively avid diggers (P. brasiliensis and A. capensis in particular) and 

groomers (all species), otters dig and groom extensively in soft loose 

natural substrates. They groom when wet or dry by rubbing, scratching, 

and digging into soft loose dry substrates, often covering their body fur with 

the freed particles. (See photo-N.A. river otter, Jennifer Potter, Calgary 

Zoo) These behaviors are among the most favored and frequently 

performed terrestrial activities in zoos and aquariums, and otters will use 

the entire expanse of their land area to carry them out. Together with 

foraging, exercising, and frequent play bouts on land, these terrestrial 

behaviors constitute a significant proportion of otters’ natural, daily goaloriented 

activities (P. brasiliensis do not forage on land). These behaviors 

are considered essential to maintaining the otters’ physical and behavioral 

health, as well as to the promotion of successful pup-rearing practices (Reed-Smith 2001; Sykes-Gatz 

2005). Additionally, the ability to carry out all of these behaviors is considered important for an otter’s 

healthy adjustment to new or unusual situations. Digging and grooming are among the most important 

activities required by P. brasiliensis in particular to prevent or reduce stress and to maintain health of the 

animals (Sykes-Gatz 2005). 

The AZA Otter SSP recommends that exhibits should be constructed of a variety of natural substrates 

to accommodate these activities. If artificial surfaces like concrete are used, these should be kept to a 

minimum. Digging pits and grooming areas with soft, loose substrates should always be included in otter 

exhibits, both indoor and outdoor enclosures. (P. brasiliensis although require additional substrate



conditions to keep sufficiently dry and healthy; see Appendix G and below in 2.1.) Adequate land area 

and substrates on which otters can groom are considered key to the care of ex-situ populations of otters. 

In order to maintain healthy thermal properties of their coats, otters have to frequently groom their fur, 

replacing the air layer trapped within the under-fur (Dunstone 1998, Weisel et al. 2005). 

A variety of live plants can be used in exhibits, as well as log piles, large tree stumps or root systems, 

hollow logs, hills, etc., all of which can provide visual complexity to the exhibit and offer otters excellent 

foraging, playing, and shelter opportunities. All of these features can be placed to allow for visitor viewing. 

However, accommodation giving very shy animals the ability to hide should be made. 

As with any species, otter exhibits should be “redecorated” periodically. However, it is advisable that 

preferred denning or hiding spots not all be changed simultaneously. All exhibits should be constructed 

with a means of accomplishing re-fitting of exhibit furniture, including the introduction of large deadfall. 

Self-dug dens, particularly those of P. brasiliensis should be allowed to remain as long as they are 

deemed safe. 

The minimum exhibit size (including land and water surface area) suggested for otters is as follows: 

• A. capensis and L. canadensis: 150m 2 (1615ft 2 ) for two animals. An extra 25m 2 (269ft 2 ) of 

useable land surface and 10m 2 (108ft 2 ) water surface should be provided for each additional 

animal (Duplaix-Hall 1975; Reed-Smith 2004a). 

• A. cinereus: 60m 2 (646ft 2 ) for 2-4 animals (Duplaix-Hall 1975); 93m 2 (1,000ft 2 ) for more than 4 

animals. 

• L. maculicollis: 100m 2 (1076.5ft 2 ) land and water surface for two animals. An extra 20m 2 (215.3ft 2 ) 

of useable land and 5m 2 (54ft 2 ) water surface should be provided for each additional animal. 

• P. brasiliensis: In North America, the minimum enclosure size (land and water surface) that has 

housed a family of giant otters is 121m 2 (1300ft 2 ). Duplaix-Hall (1972, 1975) recommends a 

minimum size of 240m 2 (2,584ft 2 ) for a pair. Sykes-Gatz (2005) also cites 240m 2 (2,584ft 2 ) for a 

pair with additional space provided for offspring. A recommended additional size for family groups 

is not offered but stress is placed in the quality of the enclosure space and land/water ratio 

appropriate for the exhibit size. Please refer to Sykes-Gatz (2005) for ratio formulas and more 

information. 

Recommended minimums are based on the species natural history, observations of in-situ 

populations of otters, and authors experience with otters in ex-situ. 

Exhibit Complexity – Aquatic: The water portion of an otter exhibit should include areas of varying 

depths, and some portion of shoreline that allows for easy access to and from the shore for both old and 

young animals. Shorelines also should be complex and designed to allow for the periodic change of 

features (e.g., logs, rock piles, pebble pockets, etc.). For more specific suggestions for P. brasiliensis 

enclosure shoreline design, see Sykes-Gatz (2005). Streambeds or shallow wading pools with rocky 

bottoms offer good enrichment and foraging opportunities, but can cause footpad abrasions if otters are 

forced to walk on them too often without access to natural substrates for drying their feet. To aid in 

minimizing the transference of debris into pool water, the shoreline can be constructed of raised 

flat/sloping rocks or logs positioned to hold natural substrates; Sykes-Gatz (2005) recommends a ~10cm 

(4-inch) curb. Furnishings should allow otters, especially pups, females carrying pups, and geriatric otters 

easy and safe ingress and egress to/from pools. Pools should be designed with several skimmers that 

can capture large particle debris before it reaches the filtration system. All pool openings such as 

skimmers, drains, filters, etc. to which the otters have access should be securely covered with sturdy wire 

fencing to prevent curious otters from getting their heads/feet stuck. Ample, extensive dry land areas 

should be provided to allow all otter species to dry off completely. 

Exhibit pools should have varying depths, offering opportunities for animals to forage in the shallow 

water and swim/dive in deeper water. Shorelines should be complex and curving, as opposed to straight 

lines and uninterrupted. Additionally, the shore should be periodically interrupted with shade structures 

(e.g., bushes, trees, etc.), and climbing or laying-out structures, such as logs, boulders, log jams, etc. 

Otters should not be forced to swim continuously next to public viewing windows as this frequently leads 

to stereotypic activities such as flip-swimming, tail-sucking, and back-flips (Reed-Smith 2004b). Instead, 

exhibit pools should offer swimming alternatives that allow the animals to access deep and shallow areas 

that are not next to the public viewing areas. A narrow pool design has not caused these negative effects 

on P. brasiliensis. Instead insufficient land vs. water area and/or inappropriate substrate conditions have 

been the primary cause of behavioral problems including; stereotypic swimming, swimming excessively



and aimlessly, and parents or older siblings overexposing pups to pools or mistreating them there (Sykes- 

Gatz 2005). See Appendix M for additional information regarding designing and maintaining pools in otter 

enclosures. 

P. brasiliensis: P. brasiliensis pools should have a deep area of at least 100cm (3.28ft), as well as shallow 

areas; the otters make frequent use of waters of differing depths. Depths of 150-200cm (5-6.5ft) and 

deeper are highly recommended allowing a wider range of swimming and diving behaviors. 

Exhibit Design and Species-appropriate Behaviors: In addition to a pool, all otter enclosures should 

be enhanced with a variety of furnishings. The quality of space for these animals is just as important as 

exhibit size. Logs, trees, tree roots, stumps, grasses, boulders, dens, caves, climbing structures, bushes, 

deadfall (positioned so animals cannot use them to climb out of the enclosure), waterfalls, floating log 

piles, rafts, islands, varied exhibit levels, and a variety of substrates are all important elements of a 

complex and successful otter exhibit (for all species). On-exhibit sleeping and hiding places should be 

provided; these sites should be of varying sizes to allow the group to sleep together or to allow for 

individual seclusion. Animals should be allowed to dig, roll, climb, and slide within their exhibit. Enclosure 

designers should take all of these activities into consideration when designing the land/water interface, 

public viewing, substrates, and pool filtration systems. Pools should be designed so that the animals are 

not always forced to swim in close proximity to the public (this requirement does not appear to be as 

crucial for the giant otter). Public viewing should be provided from various angles while maintaining one 

side without public access as a secure zone for the animals. Sykes-Gatz (2005) and Hancocks (1980) are 

two of the many resources available with information on naturalizing older exhibits. 

All pool shorelines should be provided with lounging logs, shaded rest areas, and sandy banks to be 

used as latrine sites. Large flat/sloping rocks and logs can be used along the shoreline to hold back the 

substrate as well as provide good sunning areas. 

A variety of substrates should be incorporated into otter exhibits. These substrates include: grass, 

mulch, sand, clay, soil, rocks, boulders, pebbles, leaves, bark, concrete, and gunite (the latter two are not 

recommended and should be limited to small areas, or should be covered with soft pebble-free sand or 

tree bark mulch when their use is unavoidable). The substrate recommendations for P. brasiliensis are 

slightly different to other otter species, and species-specific recommendations are provided below. 

Exhibits with artificial substrates should offer areas of grass, dirt, sand, pebbles, etc. for exploration and 

adequate grooming. Hard-pack soils, abrasive sands, and sharp rocks should not be used in otter 

exhibits. Recent research into the structure of sea otter (E. lutris) and river otter (L. canadensis) hair 

structure shows guard hairs can suffer damage (Weisel et al. 2005). While unclear at this time, it is 

possible that extensive damage to guard hairs can impact the insulative ability of the otter’s coat. 

Problems with chronically wet surface areas or overexposure to hard surfaces should be addressed 

immediately to prevent injuries to the animals (e.g., foot pad abrasions) or health issues (e.g., fungal 

infections, pneumonia) from developing. 

All exhibits should offer bedding material; products used successfully include: grasses, leaves, hay, 

straw, wood wool, sedges, pine needles, towels, burlap bags, indoor/outdoor carpeting, natural fiber 

mats, and wood shavings (Reed-Smith 2001). Bedding material recommendations for P. brasiliensis are 

different than other otter species and these species-specific recommendations are provided below. Some 

facilities have successfully used fleece and blankets (Ben-David et al. 2000, 2001a and b; J. Reed-Smith, 

personal experience). However, as with all bedding, these should be monitored to ensure the otters are 

not chewing on or eating them. If animals are chewing on these items, they should be removed 

immediately. Some wood shavings (from conifers) contain residues that can strip the water proofing from 

the coat of semi-aquatic species, and/or may cause sneezing. Cedar contains aromatic phenols that are 

irritating to the skin and respiratory system. Several studies indicated that close, chronic contact with 

cedar shavings contributed to infant mortality (Burkhart & Robinson 1978), respiratory disease (Ayars et 

al. 1989), and liver damage (Vesell 1967) in rodents. The impact of these products on otters is unknown; 

if used, caution should be exercised. 

Some facilities use paper products such as shredded paper, cardboard boxes, paper bags, and 

cardboard rolls. These products should be monitored carefully to ensure the animals are not ingesting 

them or taking them in the water where they could become plastered over an animal’s mouth and nose, 

or become impacted in their teeth. In most cases the AZA Otter SSP advises against using these 

products with otters.



Indoor/outdoor carpeting and natural fiber mats also have been used for the animals to roll and groom 

themselves on in concrete holding areas. All materials used for bedding should be monitored in case an 

animal consumes them excessively, or in the case of towels, etc., shreds or eats them. ‘Wood wool’ sticks 

to fish or other moist foods, and so should not be used near feeding areas to prevent its ingestion. When 

it is used in nest boxes, caution should be exercised if any females become pregnant as pups can 

become entangled in it. 

L. canadensis: Typically L. canadensis shed their under-fur between May and August (This “…under-fur 

produces a dense, matted, felt-like layer, which forms an efficient insulating layer by trapping air next to 

the skin…’[Dunstone 1998]), and replace their guard hair between August and November (northern 

latitudes, there may be some variation in timing at southern latitudes) (Ben-David et al. 2000; J.Reed- 

Smith, personal observation). Animals’ health requires ample grooming opportunities and surfaces on 

which they can rub to prevent matting, and aid in this annual coat replacement process. Grooming and 

drying opportunities also are important for the maintenance of healthy foot condition, with damp or 

excessively humid conditions leading to footpad abrasions. 

P. brasiliensis: Field (Duplaix 1980) and ex-situ population studies (Sykes-Gatz 2005) of P. brasiliensis 

have shown that grooming consumes a great deal of time, and plays a vital roll in group cohesion and 

home range identification. Duplaix (1980) states that this species spends fully half of its time on land with 

a great deal of this dedicated to grooming. 

Sykes-Gatz (2005) stresses the importance of providing at least the minimum land area proportion 

within the recommended land/water ratio and the recommended substrate conditions as primary to 

maintaining sufficient dry land area, nestboxes, and bedding materials in indoor and outdoor giant otter 

enclosures. The following additional designs are secondary for the same purposes, but will not be 

effective if insufficient land vs. water area or inappropriate substrate conditions (esp. hard surfaces) exist. 

The smaller the land area provided giant otter and the more land area exposed to the water’s edge, the 

harder it is to keep dry.These large otters carry significant amounts of water from their pools onto land 

during their frequent exits from the pool. Water areas should be bordered by at least 5m (16ft) of land 

area extending back from the shoreline of the pool for all exhibit sizes. For exhibits smaller than 240m² 

(2,583ft²) and up to 600m² (6,458ft²) Sykes-Gatz (2005) should be consulted for specific 

recommendations. Additionally, long water area contour lines, or land bordered by water on more than 

two sides, are not recommended for this species as this will result in the land becoming saturated as 

otters enter and exit the pool along the length of the shoreline. Dens and nest boxes should be located at 

least 3m (10ft) and preferably 5m (16ft), away from the water’s edge. Also, plentiful land area should 

extend laterally from the den/nest box entrances to offer sufficient and conveniently located areas where 

otters can dry off before entering their dens/nest boxes. These design features will help eliminate 

continuously damp or wet land surfaces and nestbox conditions, which can lead to health problems (see 

Chapter 6). 

Existing exhibits that do not meet these guidelines can be relatively easily and inexpensively 

modified. Sykes-Gatz (2005) and Hancocks (1980) provide useful information on naturalizing older 

exhibits. All enclosure surfaces can be naturalized with the methods and substrates described below. 

Sykes-Gatz (2005) stresses the importance of covering the enclosure land area and holding surface to a 

depth of 10-20cm (4-8”) with soft natural substrates (soft pebble-free sand or tree bark mulch) that are 

deep and loose enough to allow adequate drainage and remain dry, so that otters can easily dig in it, and 

adequately groom themselves upon it. The use of other mulch products is not considered advisable with 

this species. Finely pieced mulch easily becomes saturated and does not dry sufficiently, creating 

unhealthy conditions for the otters. New mulch or sand should be added on top of the existing layer as 

needed to maintain no less than the minimum recommended depth, and/or to cover broken-down or 

compacted mulch. If this is not monitored closely the substrate may become too wet or damp, or too hard 

and unusable for digging or grooming by the otters. Additionally, indoor and outdoor enclosures should 

each have an area (20m² (215ft²) is recommended by Sykes-Gatz, 2005) with sand or mulch at least 40- 

60cm (16-24") deep, or soil hillsides, to allow for deep digging. See Appendix G and Sykes-Gatz (2005) 

for additional information on appropriate substrates and depths for giant otters. 

Animal shift and keeper doorframes raised roughly 10cm (4") above the desirable substrate surface 

height will help prevent substrates from blocking all doors. A log border placed a little behind an existing 

keeper door or placed against the front and back of an animal shift doorframe, or a wooden lip for the 

door’s track, serves the same purpose (Sykes-Gatz 2005).



Nest boxes can be provided with sand or mulch that is 10cm (4") in depth. When young pups are not 

present, other bedding materials such as wood wool, hay, straw, or leaves also may be provided, but 

these materials should be removed prior to birth of any litters and can be replaced with sand or tree bark 

mulch. Care should be exercised to ensure that all nest box substrates remain dry. Damp conditions may 

contribute to otter pup mortality. After parturition, otters may dig out all bedding but can still successfully 

raise pups. Pine needles, towels, burlap bags, indoor/outdoor carpeting, and natural fiber mats should not 

be offered to this species (Sykes-Gatz 2005). 

The recommended substrates are inexpensive, effective, easy to maintain and acquire, and they 

remain sanitary with dry spot cleaning. When more land area is needed for an appropriate land/water 

ratio, a portion of an artificial (e.g., concrete) or natural pool can be divided with a waterproof barrier, or 

one or more of multiple pools can be emptied. These can be filled-in with the recommended substrates to 

create enough land. 

This species in particular appears to like digging underneath large tree stumps with long root 

systems, logs, etc., and so these furnishings are advisable, especially in deep digging areas. A variety of 

vegetation and furnishings of this type, in addition to grasses, bamboo stands, and leaf piles, should be 

contoured into all exhibits allowing for exploration, visual barriers, and privacy when pups are born. Other 

effective furnishings include large hollow logs, large logs on land and lying into-over the water, and 

boulders (Sykes-Gatz 2005). In climates where otters can be outside year around, some zoos have 

successfully provided soil hillsides to allow otters to dig underground dens. The hillsides should be at 

least 2m (6.5ft) high, have an angle not more or less than 40-45°, and be located behind and near the 

water area shoreline (otters cannot dig deep enough in flat terrain or shallow substrates) (Sykes-Gatz 

2005). Trees, large bushes, or tree stumps with long extended root systems may help prevent den caveins 

but this is always a danger. If there is any doubt about the safety of a den it should be refilled with 

substrate. See Chapter 2 for enclosure barrier considerations to prevent escape by digging. See 

Appendix G for additional information on giant otter exhibit furnishings, substrates, etc. 

Sensory Barriers: Visual barriers are important to allow animals to avoid one another, when necessary. 

All individuals, particularly paired otters, will go through times when they exhibit a tendency to stay by 

themselves. Vegetation, exhibit topography, denning sites, and deadfall should be strategically placed to 

allow for this. While there is no evidence that low level background noise is disruptive to otters, loud 

noises can be frightening to them, and high-pitched, long-term noise should be avoided. 

Otters can be odiferous; facilities with indoor exhibits may want to provide olfactory barriers for the 

comfort of the viewing public. If exhibiting more than one breeding otter group, or permanently separating 

animals from a group and housing them within the same institution, it may be very important to have 

visual, olfactory, and audio separation to avoid intra-specific aggression or abnormally elevated levels of 

stress and/or frustration. In these cases, it is advisable to plan for this in advance rather than trying to 

accommodate it if a problem arises. 

L. canadensis: Parturient females generally become very aggressive towards the male several days 

before giving birth and while the pups are quite young. Pairs housed in large naturalistic exhibits can be 

maintained together if sufficient visual barriers are provided to allow the male to remain out of the 

female’s line of sight. In all other cases, it is important that the pair is separated. The pair can be left at 

the exhibit if one animal is held in the holding dens/off-exhibit area while the other is on exhibit. In these 

instances, the animals should not have to pass in view of one another to shift into alternate areas for 

cleaning or feeding. 

P. brasiliensis: Visual and acoustic isolation from human disturbances (staff and visitors) is necessary 

during parturition and early pup-rearing. All human sounds and disturbances should be minimized to 

ensure successful pup rearing, as this species is highly sensitive to human interference. See Chapter 7 

for further information. 

In the rare cases when bonded pairs or other group members have to be separated, they should be 

held in facilities distant enough to prevent visual, acoustic, or olfactory communication. P. brasiliensis is 

highly vocal and their calls carry great distances. 

Enclosure Cleaning: Otters are scent-oriented animals; therefore, their entire exhibit or holding area 

should not be cleaned at the same time. Enclosures should be raked and spot cleaned daily, with 

appropriate disinfecting as necessary. Indoor or hard surface floors should be cleaned with detergent 

daily. Due to their natural scent marking behavior, exhibit furniture should not be cleaned as frequently. It



may actually be stressful to some otters if their territory is totally cleared of their markings. All detergents 

should be thoroughly rinsed, as any residue left on the floor, mats, or furnishings can strip their coats of 

its natural oils (Schollhamer 1987). 

Food and water containers should be cleaned and disinfected daily. All food remains should be 

removed before it can become spoiled; in some climates this 

may require removal more than once a day. A safe and effective 

control program for insects, ectoparasites, and bird and mammal 

pests also should be maintained. 

The same careful consideration regarding exhibit size and 

complexity and its relationship to the otter’s overall well-being 

must be given to the design and size all enclosures, including 

those used in exhibits, holding areas, hospital, and 

quarantine/isolation (10.3.3). 

A holding area connected to the exhibit is recommended; 

ideally this should include a pool with clean water available at all times, proper lighting, appropriate 

lighting cycle, a sleeping or den box, enough floor space for grooming and drying areas, an outdoor play 

area (if extended stays are routine), and at least one nest box that is heavily bedded to allow excess 

moisture to be removed from the animals’ coats (Lombardi et al. 1998; Reed-Smith 2001). Each holding 

den should be large enough to at a minimum allow the animal to turn around (when used only for holding 

during cleaning) but at least 122cm (4 ft.) by 122cm (4 ft.) if an otter is to be held over-night. Best 

practices would include larger night dens than this minimum. Holding pens should have non-climbable 

sides, access one to another (allowing animals to have more room during extended stays), and multiple 

ingress and egress to the exhibit. Maternity dens should be isolated so exhibit mates do not have to pass 

in front of this den to enter or exit the exhibit. If chain-link barriers are present, the sides should be 

covered with lexan or similar material to prevent animals from climbing too high and falling. 

Quarantine facilities should offer the maximum space available. They should be furnished with natural 

items to include logs, stumps, and outdoor carpeting that can be periodically disinfected/changed as well 

as sand/mulch/straw/soil boxes that can be changed when they become soiled or saturated. 

2.2 Safety and Containment 

Otters are not appropriate for free-ranging environments. 

Animal exhibits and holding areas in all AZA-accredited 

institutions must be secured to prevent unintentional animal 

egress (11.3.1). Exhibit design should be considered carefully to 

ensure that all areas are secure and particular attention must be 

given to shift doors, gates, keeper access doors, locking 

mechanisms and exhibit barrier dimensions and construction. 

Otters can climb, and will take advantage of anything 

provided. Trees, bushes, etc., should be placed away from exhibit 

perimeters. Walls should be non-climbable, and fences should be 

strong and inhibit climbing. The containment walls/fence should 

be at least 1.52m (5ft) high for A. cinereus and at least 1.83m 

(6ft) high for L. canadensis, L. maculicollis, P. brasiliensis, and A. 

capensis. While these heights should contain most otters, Ben- 

David (personal communication) reported a L. canadensis scaling 

a 3m (9.8ft) fence. Animals that are known to be climbers may 

require additional containment height or features. It has been 

shown that Lutra lutra can clear 1.3m (4.27ft) when leaping from 

the ground to a platform, 1.6m (5.25ft) when jumping from one 

platform to another, and 0.92m (3ft) when jumping from the water 

onto a platform, if they are able to push off from the pool bottom 

(Reuther 1991). If containment barriers are mesh, they should be 

topped with an un-climbable, inward-facing overhang of 80cm 


(10.3.3) All animal enclosures (exhibits, 

holding areas, hospital, and 

quarantine/isolation) must be of a size 

and complexity sufficient to provide for 

the animal’s physical, social, and 

psychological well-being; and exhibit 

enclosures must include provisions for the 

behavioral enrichment of the animals. 


(11.3.1) All animal exhibits and holding 

areas must be secured to prevent 

unintentional animal egress. 


(11.3.6) Guardrails/barriers must be 

constructed in all areas where the visiting 

public could have contact with other than 

handleable animals. 


(11.2.3) All emergency procedures must 

be written and provided to staff and, 

where appropriate, to volunteers. 

Appropriate emergency procedures must 

be readily available for reference in the 

event of an actual emergency. These 

procedures should deal with four basic 

types of emergencies: fire, 

weather/environment; injury to staff or a 

visitor; animal escape. 

(2.7ft) (Duplaix-Hall 1975; Foster-Turley 1990). Hot wire can be used, but should not be accessible to an 

animal in the water, and should be placed at a height that will not cause injury to an animal if they fall as a 

result of touching the wire.



Otters also dig proficiently (particularly A. capensis, L. canadensis, and P. brasiliensis). When 

designing exhibits, perimeter walls and fences should be buried or mesh linked. Sinking perimeter 

fences/walls at least 80cm (2.6ft) is advisable for most species, however, Sykes-Gatz (2005) 

recommends sinking perimeter fences deeper than 1m (3.28ft) for P. brasiliensis, because this species 

easily digs down that far. Holes along perimeter containment should be promptly refilled. To provide 

additional safety, secondary containment areas should be constructed at all enclosure entrances. 

Exhibits in which the visiting public may have contact with animals must have a guardrail/barrier that 

separates the two (11.3.6). Barriers should be high enough to prevent visitors from reaching over or into 

the exhibit. 

In case of an otter escaping institutional and AZA policies for containment should be followed. 

Typically all otters will run and hide if they are not able to return to their exhibit. Staff should notify the 

appropriate chain of command, keep the animal in sight, ask the public to leave the area, and attempt to 

slowly encourage the animal into a building or other confined area. Nets, push boards, and gloves should 

be availbe in case of these emergencies. Tranquilizer guns should not be used on these species, 

particularly the smaller ones. Instead, if anesthesia is required a 

blow pipe should be used. In the case of severe weather (i.e. 

hurricanes, tornados, thunderstorms animals should be confined 

to indoor holding. Policies regarding other emergencies such as 

fire or earthquake should be developed by each institution based 

on the species and enclosure design. For all of these situations, 

an institutional policy should be developed in advance and all 

emergency safety procedures should be clearly written, provided 

to appropriate staff and volunteers, and readily available for 

reference in the event of an actual emergency (11.2.3). 

Staff training for emergencies must be undertaken and 

records of such training maintained. Security personnel must be 

trained to handle all emergencies in full accordance with the 

policies and procedures of the institution and in some cases, may 

be in charge of the respective emergency (11.6.2). 

Emergency drills should be conducted at least once annually 

for each basic type of emergency to ensure all staff is aware of 

emergency procedures and to identify potential problematic areas 

that may require adjustment. These drills should be recorded and 

evaluated to ensure that procedures are being followed, that staff 

training is effective and that what is learned is used to correct 

and/or improve the emergency procedures. Records of these 

drills should be maintained and improvements in the procedures 

duly noted whenever such are identified. AZA-accredited 

institutions must have a communication system that can be 

quickly accessed in case of an emergency (11.2.4). 

AZA-accredited institutions must also ensure that written 

protocols define how and when local police or other emergency 

agencies are contacted and specify response times to 

emergencies (11.2.5). AZA-accredited institutions which care for 

potentially dangerous animals must have appropriate safety 

procedures in place to prevent attacks and injuries by these 

animals (11.5.3). Animal attack emergency response procedures 

must be defined and personnel must be trained for these 

protocols (11.5.3). 

Otter attack emergency drills should be conducted at least 

once annually to ensure that the institution’s staff know their 

duties and responsibilities and know how to handle emergencies 

properly when they occur. All drills need to be recorded and 

evaluated to ensure that procedures are being followed, that staff 

training is effective, and that what is learned is used to correct 

and/or improve the emergency procedures. Records of these 


(11.6.2) Security personnel, whether staff 

of the institution, or a provided and/or 

contracted service, must be trained to 

handle all emergencies in full accordance 

with the policies and procedures of the 

institution. In some cases, it is recognized 

that Security personnel may be in charge 

of the respective emergency (i.e., 

shooting teams). 


(11.2.4) The institution must have a 

communication system that can be 

quickly accessed in case of an 

emergency. 


(11.2.5) A written protocol should be 

developed involving local police or other 

emergency agencies and include 

response times to emergencies. 


(11.5.3) Institutions maintaining 

potentially dangerous animals (sharks, 

whales, tigers, bears, etc.) must have 

appropriate safety procedures in place to 

prevent attacks and injuries by these 

animals. Appropriate response 

procedures must also be in place to deal 

with an attack resulting in an injury. These 

procedures must be practiced routinely 

per the emergency drill requirements 

contained in these standards. Whenever 

injuries result from these incidents, a 

written account outlining the cause of the 

incident, how the injury was handled, and 

a description of any resulting changes to 

either the safety procedures or the 

physical facility must be prepared and 

maintained for five years from the date of 

the incident.



drills must be maintained and improvements in the procedures duly noted whenever such are identified 

(11.5.3). 

If an otter attack occurs and injuries result from the incident, a written account outlining the cause of 

the incident, how the injury was handled, and a description of any resulting changes to either the safety 

procedures or the physical facility must be prepared and maintained for five years from the date of the 

incident (11.5.3).

3.1 Preparations 

Animal transportation must be conducted in a manner that 

adheres to all laws, is safe, and minimizes risk to the animal(s), 

employees, and general public (1.5.11). Safe otter transport 

requires the use of appropriate conveyance and equipment that is 

in good working order. 

When transporting otters there always should be at least two 

people present; if animals have been anesthetized the veterinarian 

always should be present. Staff involved in transports should 



Chapter 3. Transport 


(1.5.11) Animal transportation must be 

conducted in a manner that is safe, 

well-planned and coordinated, and 

minimizes risk to the animal(s), 

employees, and general public. All 

applicable local, state, and federal 

laws must be adhered to. 

understand their duties and have a clear idea of the institutions policies regarding transports. The Otter 

SSP has no specific recommendations regarding staff roles in transports but does recommend 

procedures and policies be clearly defined and understood in advance by all participating staff. 

The equipment must provide for the adequate containment, life support, comfort, temperature control, 

food/water, and safety of the otters(s). Safe transport also requires the assignment of an adequate 

number of appropriately trained personnel (by institution or contractor) who are equipped and prepared to 

handle contingencies and/or emergencies that may occur in the course of transport. Planning and 

coordination for animal transport requires good communication among all affected parties, plans for a 

variety of emergencies and contingencies that may arise, and timely execution of the transport. At no time 

should the otter(s) or people be subjected to unnecessary risk or danger. 

The transport of wild animals is regulated by the International Air Transport Association (IATA). The 

standards of care provided within this chapter are based on IATA regulations (e.g., IATA 2007), best 

practice recommendations from the AZA Otter SSP, and AZA Accreditation Standards. 

Pre-shipment Exams: All otters should receive a thorough pre-shipment physical examination (for more 

information, see Chapter 6). Ideally, a copy of the pre-shipment physical exam findings and laboratory 

work should be sent to the veterinarian at the receiving institution before the animal is transferred. If an 

otter has a current medical condition requiring ongoing treatment, the case should be discussed between 

the veterinarians at the shipping and receiving institutions before the animal is moved. All animal 

shipments should be accompanied by a hard copy of the medical record, as well as a health certificate 

and the USDA acquisition, disposition, or transport form (form #7020) (APHIS 1997) as well as any other 

paperwork required by IATA, carrier, or regulatory agency. Institutions using MedARKS should provide 

the receiving institution with electronic copies of the medical records. Dietary, enrichment, and training 

records should be sent prior to shipping the animal 

Crate Requirements: All possible relevant regulatory agencies should be checked for shipping, health, 

and permit requirements before transporting animals (USFW, state regulations, CITES, etc.). The 

International Air Transport Association (IATA) publishes specific guidelines for transport containers used 

for animal shipments. These guidelines are available from the Publication Assistant, IATA, 2000 Peel 

Street, Montreal, Quebec, Canada, H3A 2R4 (Ott Joslin & Collins 1999). An alternate address for IATA is 

International Air Transport Association, 800 Place Victoria, P.O. Box 113, Montreal, Quebec, Canada, 

H4Z 1M1. The Live Animals Regulations document is available in print or CD ROM format, and can be 

ordered from sales@iata.org. IATA regulations change periodically, and so the most recent publication or 

website should be consulted. It is very important to adhere closely to these requirements as airlines may 

refuse to fly animals in containers that do not conform to the guidelines. In general, IATA regulations 

require the following: 

All shipping crates should allow for adequate ventilation. Ventilation apertures should be small 

enough to prevent the escape of the animal and small enough that the animal cannot get any part of its 

body through the opening. 

Generally, a Vari Kennel ® may be used for A. capensis, A. cinereus, L. canadensis, and L. 

maculicollis, with the following modifications. The grill door should be covered with securely fixed weld 

mesh and all ventilation openings covered with wire mesh. The door should have secure fasteners at the 

top and the bottom. A curtain that can be raised and lowered and does not block ventilation should be 

fixed over the door to reduce light inside the container. A dropping tray should be fixed to the floor and



filled with absorbent material; the container should be leak-proof. There should be ventilation openings on 

the rear of the container; extra ventilation openings may have to be added in order that the total 

ventilation area is at least 20% of the four sides. The container should be correctly labeled. If the 

container has wheels, they should be removed or rendered inoperable. Airlines also may require a wire 

mesh cage be fitted to the inside of the Vari Kennel ® . 

Shipping crate doors should be secured with additional fasteners; A. cinereus has been known to pull 

doors inwards when they could not push them out and escape from containers that were not fastened 

securely. This proviso applies to all otter species. 

Otters should be transported separately. A. cinereus have been transported successfully in groups 

when the animals are less than six months of age. IATA states that “the height of the container must allow 

the animal to stand in a natural position with its head extended and the width must permit it to turn around 

and lie down comfortably” (IATA 2009). Sykes-Gatz (2005) reports successful shipments of P. brasiliensis 

in containers measuring 140cm (55.2in) in length, 60cm (23.6in) in width, and 57cm (22.5in) tall. 

However, all IATA and airline regulations should be checked prior to selecting or constructing shipping 

containers. 

P. brasiliensis: Sykes-Gatz (2005) states that P. brasiliensis should not be transported in hard plastic 

containers in situations other than within an institution. Airlines and IATA regulations should be consulted 

for appropriate transport containers for this species. It is advisable to ship this species without a wire 

mesh lining, as it could be harmful to their exceptionally sensitive footpads and webbing between the toes 

(Sykes-Gatz 2005). Lining of this type has not been proven necessary for safety when transporting this 

species. However, IATA requires that giant otter be shipped in containers with an interior of heavy-gauge 

wire mesh or sheet metal lining (IATA 2009). IATA regulations require the floor to be solid-metal, leak 

proof and covered with a thick layer of absorbent material (e.g. shavings). It is very important that the 

relevant IATA and airline regulations be checked prior to constructing all shipping containers and 

complied with to ensure acceptance by the transport carrier. 

3.2 Protocols 

Transport protocols should be well defined and clear to all otter care staff. Institutional protocols for 

planning and carrying out transports should be used for otters. There are no specific requirements. 

However, the AZA Otter SSP does encourage facilities to train their animals to voluntarily enter crates for 

transport. 

Food and Water: 

IATA requires that the crate allows for feeding and watering of the animal if needed. The food and 

water ports should be clearly marked on the outside of the crate. In case of delay on long flights, 

provisions should be made for feeding in transit (this may necessitate shipping food with the animal). 

Otters do not normally require additional food and water during transport for periods up to 24 hours. If 

unforeseen delays occur, canned dog food or cat food may be offered. Adequate moisture is present in 

these foods to take care of water needs during short periods. A metal bowl may be attached to the corner 

of the crate; this should be accessible from the outside. Always check for any specific IATA or shipping 

carrier regulations or requirements. 

Bedding and Substrate: Bedding, such as straw or shavings, should be placed in the transport container 

for the animal’s comfort and absorption of feces and urine. Types of bedding material allowed should be 

checked with the airline. In order to separate urine and feces from the animal, IATA requires a drop tray 

be fixed to the floor of the crate and filled with absorbent material. 

Temperature: Temperatures of 7.2-26.7°C (45-80°F) that are permitted by the airlines for transport are 

not appropriate at the high end for otter species. Otters should not be exposed to 21.1°C (70°F) 

temperatures for periods longer than 15 minutes when contained in a shipping crate. These animals can 

easily overheat at elevated temperatures (21.1°C or 70°F and higher), especially when stressed and/or 

contained in a shipping crate. While this is a more restrictive temperature range than previously 

recommended, past experience has shown that otters quickly overheat and succumb to hyperthermia. 

Animals in unheated vehicles should not be exposed to temperatures less than 7.2°C (45°F). 

A. cinereus: A. cinereus should not be exposed to drafty 7.2°C (45°F) temperatures for extended periods. 

This also may be true for L. maculicollis, but it is not known at this time and requires further research.



L. canadensis: L. canadensis exhibit a low heat tolerance and should not be shipped when temperatures 

are forecasted to exceed 21.1°C (70°F) at transfer locations. 

P. brasiliensis: P. brasiliensis have been shown to have low heat tolerance. It is recommended that this 

species should not be transported in temperatures below 15.5°C (60°F) or above 26.6°C (80°F) (Sykes- 

Gatz 2005). 

Light and Sound: As far as possible, noise should be kept to a minimum (including sudden loud noises, 

constant high-pitch noises, or anything considered uncomfortable to people), and the animal kept in low 

light conditions. Mesh doors or side windows (i.e., as in air kennels) should be covered with a breathable, 

opaque material to allow for ventilation and privacy for the animal (Ott Joslin & Collins 1999). These basic 

noise reduction precautions will help reduce stress from sudden, frightening sounds, and low light will 

provide a minimal sense of security for species that hide in small, dark spaces (e.g., dens) when 

frightened. 

Animal Monitoring: Otters should not be maintained in shipping containers for longer than 24 hours 

without food and water. Directions on what should be done in the case of an emergency that necessitates 

animal treatment should accompany all animal shipments. If access to an animal is required due to illness 

or extended shipment delays, the shipping/receiving institutions should be notified. The crate should be 

transferred to the nearest zoo or veterinary clinic prepared to handle the animal. 

Emergencies: In case of emergencies contact information for the shipping and recipient zoos should be 

included on the paper work and the crate. If an otter escapes, the animals should be closed into a 

confined space (if possible) and both institutions notified as quickly as possible. At a minimum, visual 

contact (from a distance) should be maintained until one or both institutions have been contacted and 

professionals have arrived. 

Post-transport Release: Upon arrival at their destination, shipping crates should be placed inside the 

quarantine holding pen, the door opened, and the animal left to exit at will. All holding pens should be 

provided with food, water, alternate hiding places, appropriate bedding, and enrichment structures.



Chapter 4. Social Environment 

4.1 Group Structure and Size 

Careful consideration should be given to ensure that otter group structures and sizes meet the social, 

physical, and psychological well-being of those animals and facilitate species-appropriate behaviors. 

A. cinereus: In zoos and aquariums, Asian small-clawed otters are monogamous, with both members of 

the pair helping to raise the offspring. Unlike L. canadensis, the otter parents and offspring should be 

housed together. Older siblings help raise the younger ones, and a family group or breeding pair can 

produce two litters a year with up to seven pups in each litter. Similar sized groups are sometimes found 

in the wild (Foster-Turley 1990). It is recommended that these otters be held in adult pairs, adult pairs 

with offspring, or single sex groups. The size of the group will depend on the size of the exhibit and the 

compatibility of the individual animals (Lombardi et al. 1998). The formation of single sex groups should 

be accomplished at a very young age to avoid aggression. 

A. capensis: A. capensis have been observed in multiple social situations. In fresh water inland systems, 

the male stays with the female and offspring (Rowe-Rowe 1978). Clans up to 10-15 individuals have been 

observed. In marine ecosystems, it has been noted that the male does not stay with the family group 

(Estes 1989). Females with pups are the typical family group (Chanin 1985), but this species is often 

seen alone (Chanin 1985). In zoos and aquariums, pairs have been separated during parturition and early 

pup rearing. This species has been held in pairs and in groups of 1.2 in zoos and aquariums (Reed-Smith 

& Polechla 2002). 

L. canadensis: L. canadensis are believed to be more social than most other mustelids (but not as social 

as some of the other otters), based on the findings of a number of researchers (Beckel 1982; Rock et al. 

1994; Testa et al. 1994; Johnson & Berkley 1999; Blundell et al. 2002a,b; Gorman et al. 2006, M. Ben- 

David, personal communication; S. Shannon, personal communication). For example, Stevens & Serfass 

(2008) documented that out of 172 film documentations of otters using latrines, 59.3% were by single 

otters, 19.2% were by two otters, 17.4% were by three otters, and 4.1% were by four otters. 

A variety of social groupings have been documented, but in general, the following are most typical: 

female with offspring; lone male; group of males; lone female; group of males with sub-adult females; pair 

(during mating season only); or two females with offspring. Male groups have been reported by Blundell 

et al. (2002a, 2002b, 2004) and Hansen (2004); female groups were recorded by Gorman et al. (2006) 

Otter associations may vary with the habitat in which the animals are found. Blundell et al. (2002a,b) 

found otters living in areas rich in resources seem to show more of a tendency to socialize, particularly 

the males and sub-adult females. Discounting female and pup associations, the Blundell et al. (2002a,b) 

telemetry study found that females were asocial in 47% of their locations, while males were asocial during 

only 24% of their locations. Further, they determined that among the “social” otters, males were social in 

46% of their locations and 63% of that time was spent in all male groupings. “Social” females were 

located in social groupings only 26% of the time, and 78% of that time they were located in mixed-sex 

groupings (Blundell et al. 2002a,b). 

Gorman et al. (2006) found in Minnesota that annual home ranges for males were 3.2 times greater 

than for females, and annual core areas for males were 2.9 times greater than for females. Core areas 

frequently overlapped with interactions between overlapping resident females and males most common 

(51.7%), while male-male (27.6%) and female-female (20.7%) interactions were slightly less frequent. 

Female core areas overlapped those of other females 22.2% of the time and males 15.8% of the time; 

male core areas overlapped with those of other males on average 15.7% of the time, on average. They 

concluded that river otters living in the upper Mississippi watershed “exhibited clear evidence of space 

sharing, suggesting that individuals in this population were neither solitary nor territorial….[otters] 

appeared to socialize to some degree with any individual for which they had an encounter opportunity.” 

Zoos and aquariums would qualify as “habitats rich in resources,” thus providing the opportunity to 

keep more social groupings of this species as found in some portions of their range. Ben-David et al. 

(2000, 2001a, and b) maintained 15 unrelated males in one enclosure for 10 months with little to no 

problem. 

There are no minimum or optimum group sizes for this species. To the contrary, this species’ 

behavioral plasticity allows the formation of social groups not normally associated with a typically solitary 

species. Preferred groupings include: multiple males, a male-female pair, one male and multiple females



(1.2), multiple pairs (2.2). The only social grouping not recommended is all female, unless they are 

related (or introduced at a very young age) and have been together continuously (Reed-Smith 2001). This 

does not mean it is impossible to introduce adult females or house unrelated females together, just that it 

is very difficult and frequently unsuccessful (Reed-Smith 2001, 2004b). See Introductions/Reintroductions 

and Appendix N for additional information. There are some indications that pairs raised together do not 

breed well; in situations where breeding is desired, one of these young otters may need to be switched 

with an unfamiliar animal. 

Multiple pair groupings should be monitored closely for signs of stress in subordinate animals. 

Groupings of multiple males with one female are not recommended, but can be maintained if monitored 

closely or separated during breeding season to prevent the males from fighting over the female or 

causing harm to the female with overly attentive advances (Reed-Smith 2001). 

L. maculicollis: This species is best housed as pairs or family groups. All male groups also may be a good 

option but not enough is known yet to determine how this natural social pattern works in a confined 

setting. While several field researchers have reported seeing large groups (10-20 individuals) of L. 

maculicollis (Proctor 1963; Kruuk & Moorhouse 1990; Kruuk 1995, J. Reed-Smith in prep), it is not known 

what role, if any, older siblings play in caring for younger pups or how often family groups join. Proctor 

(1963) reported observing groups of about five otters most frequently, a size believed to be consistent 

with a single family. Ongoing studies in Lake Victoria cite frequent observation of groups of 6-8 animals, 

at times constituted of animals of varying sizes (J.Reed-Smith, personal observation). 

It is considered unwise to introduce adult males and difficult to introduce adult females. In the latter 

case, some females show a marked preference for certain females over others. Compatible groups of 

females may show aggression towards one another at times, particularly if a female is in estrus 

(Scollhamer 1987). In one instance, older pups have been maintained in an exhibit with the adult pair and 

younger pups (R.Willison, personal communication); the female kept the older pup and adult male at bay 

as parturition neared. However, there have been reports of adult females ostracizing female offspring 

which required the young female to be moved (Benza et al. 2009). Reports from the wild that this species 

typically live as pairs, with both parents participating in pup rearing, have come under some doubt (Reed- 

Smith in prep). This has not been the case in zoos and aquariums, at least during the early phase of pup 

rearing; instead, the female raises the pups alone until they are active and swimming at which time the 

male is again allowed into the group (R.Willison, personal communication). Recent field studies in Lake 

Victoria, Tanzania (Reed-Smith in prep) recorded single animals, mother with young, two or more adult 

females with young, male groups/pairs and adolescent groups as the social configurations sighted. 

Optimal group size will vary with the size of the exhibit and compatibility of individuals involved, as 

with all otter species. 

P. brasiliensis: There is a high degree of pair bonding and group cohesiveness in Pteronura (Duplaix 

1980). In the wild, a mated pair normally bonds for life, and all family members, including offspring (one 

and two year olds) from previous litters, care for new pups (Schenck & Staib 1994). In zoos and 

aquariums, this species should be housed as mated pairs with young up to about two years of age. All 

male groups (e.g., 2-3 animals) can be exhibited as an alternative. Females can be kept together, but 

these are generally only successful as related duos or animals introduced at an early age. There are a 

few reports of adult females living together ex-situ, but it is not known if these animals were related. There 

are no reports of the successful introduction of adult females and this grouping is not recommended. 

There are some indications that pairs reared together from a very young age, or introduced well 

before they reach sexual maturity, will not breed successfully (Sykes-Gatz 2005). Therefore, it is 

recommended that breeding pairs be introduced after they have matured sexually. 

In the wild, one litter of 1-6 pups is born annually. The pups are dependent upon the other family 

members for care, socialization, learning life skills, etc. In zoos and aquariums, the pair should be allowed 

to rear their pups together. Preventing such activities often causes significant problems, such as litter 

loss, and inhibits the socialization of older young who have to learn pup care from their parents. Both 

parents will take care of the pups, teach them how to swim, feed them, groom them, etc. On occasion, 

both will move them to different nest boxes. In the wild (Staib 2002) and in zoos and aquariums (Sykes- 

Gatz 2005), the sire may sometimes take pups from the den only to have them immediately returned 

there by the dam. This behavior is considered normal and should not be viewed with alarm unless it is 

accompanied by signs of agitation or excessive stress (Sykes-Gatz 2005). See Chapter 7 for additional 

information on pup rearing in this species.



Single-sexed Groups: The following recommendations are provided for the formation and maintenance 

of single-sexed otter groups: 

A. cinereus: Single-sexed groups are not seen in the wild, but all male groups are found in zoos and 

aquariums. Groups should be formed as early and at as young an age as possible; extra care should be 

given at feeding time. Extra dens should be provided so that every animal has its own area to be apart 

from the others (Lombardi et al. 1998). 

While it is possible to keep single-sex sibling groups, it is not a preferred grouping and should be 

attempted only at the recommendation of the AZA Otter SSP. Generally, the group structure begins to 

break down beginning at sexual maturity, leading to increasing aggression toward subordinate animals. 

Typically, all male groups have been more successful than female groups and hinge on introduction of 

non-sibling animals at a very young age. 

In one case, six females housed together all of their lives began fighting at the ages of 3-5 years, 

when the dominant female began harassing the most submissive animal. This behavior was not 

apparently associated with estrus, and reduction of the aggression was originally handled through training 

(the animals had been conditioned to separate into two groups and station singly for brief periods) and 

positive reinforcement (e.g., the aggressive and subordinate animals were reinforced for looking calmly at 

one another – this was not successful). However, with time, the aggressive behavior focused on expelling 

the subordinate female spread throughout the group and previously friendly otters began to have 

problems as well. The situation eventually resulted in the need to permanently separate the dominant and 

subordinate animals, at first rotating them in with different group members and then sending them out to 

different institutions (S.Duncan, personal communication). 

A. capensis: Information on the success of single-sexed groups is unknown at this time, and further 

research is required. 

L. canadensis: All male groups do very well together and have been repeatedly documented in the wild, 

particularly in resource rich environments (Blundell et al. 2002a and b; G.Blundell, personal 

communication; S.Shannon, personal communication, Gorman et al. 2006). Fifteen unrelated adult males 

were successfully housed together for 10 months at the Alaska Sea Life Center (Ben-David et al. 2000, 

2001a,b), and five males have been successfully housed together at the Virginia Aquarium (C.Harshaw, 

personal communication). All male groupings are suggested for non-breeding situations. 

Sub-adult females (older than one year but less than two years of age or prior to first estrus) and 

females without pups (this occurs less often) have been known to associate with all male groups (Blundell 

et al. 2004). All female groups generally do not fare well, however these groups have been reported in the 

wild by Gorman et al. (2006). In zoos and aquariums, sisters or females introduced at an early age may 

be compatible for years. However, if one animal has to be separated, they may not re-establish this social 

equilibrium upon reintroduction (Reed-Smith 2001, see Appendix N for introduction/reintroduction 

protocol). In one other in-situ study, an all female clan was identified (S.Shannon, personal 

communication). All female groupings are not generally recommended. 

L. maculicollis: Adult males introduced before the age of four months have been kept together 

successfully, as long as they are not exposed to the scent or sight of estrus females (Schollhamer 1987). 

Adult females can be housed together, but their introduction may be difficult. Some females show a 

marked preference for certain females and a dislike of others. Females that have been previously housed 

together can be reintroduced after giving birth, once their pups are eating solid food at about three 

months of age (Schollhamer 1987). Aggression has been shown towards younger females but 

documented only in mixed-sex situations (Benza et al. 2009). Further research is required to determine 

the most compatible social groupings for this species. 

P. brasiliensis: Single sex groups, typically two males, have been kept together successfully (rarely have 

groupings of three males been successful). However, these groupings should be raised together or 

introduced slowly to monitor any signs of potential aggression. There are a few reports of adult females 

living together but no additional information is available. The introduction of adult females to one another 

is not advised (Sykes-Gatz 2005).



4.2 Influence of Other Species and Conspecifics 

Compatible otters typically do not require specific inter-individual distances. All animals should be 

provided with denning choices, allowing them the opportunity to sleep together or separately. Breeding 

pairs of A. capensis, L. canadensis, and L. maculicollis will require separation during the early stages of 

pup rearing or an exhibit environment that allows for physical and visual separation of the male and 

female. 

While it is not recommended to house Asian small-clawed otter groups within visual or auditory range 

of each other (Lombardi et al. 1998), different groups of spotted-necked otter (R.Willison, personal 

communication), Eurasian otter (cited in Reed-Smith 2004b), and North American otters (Reed-Smith 

2004b) have been successfully housed within auditory and visual range of one another. Care always 

should be taken that these groups are not experiencing undue stress; all animals should be monitored for 

any signs of stress or agitation (such as excessive grooming, screaming, pacing, or squabbling through 

containment wire). When animals are separated from a group for any reason (all species), care should be 

taken when reintroducing them; at times, all of the steps of an initial introduction should be followed (see 

4.3 Introductions and Reintroductions). Some of these reintroductions, particularly with females, may not 

be successful, so prolonged separation, particularly of Asian small-clawed otters and giant otters, should 

be avoided except in cases of medical treatment, aggression, or parturition (this last condition is not 

applicable to Asian small-clawed and giant otters because they should be maintained within their groups 

for parturition). 

A. cinereus: If a group of Asian small-clawed otters have to be split for some reason (e.g., aggression 

between two individuals), experience has shown that the non-aggressing animals should be rotated 

between the two groups to prevent aggression between additional animals. In these cases, the social 

bond between highly compatible animals appeared to break down if the animals were left separated from 

any particular individuals for more than about two weeks. These temporary groups were rotated between 

off-exhibit holding and the exhibit on a daily basis, allowing all animals time in each location, but 

essentially keeping the problem animals separated visually and physically (S.Duncan, personal 


Mixed-species Exhibits: All individuals housed in mixed-species exhibits should be routinely monitored 

for stress, injuries, and to ensure they are getting adequate food and water. Typically, otters are exhibited 

with species that focus behaviorally on other exhibit features than those used by the otters (e.g., arboreal 

species). Animals within mixed exhibits should be monitored for stress, and management plans should be 

made to accommodate older animals, special nutritional needs, impending births, etc. 

A. cinereus: This species has been exhibited successfully with barbirusa, binturong, black hornbills, 

butterflies, peafowl, gibbons, giant hornbill, muntjac, Prevost squirrels, proboscis monkeys, slender-nose 

crocodiles, giant Asian squirrels, and Rodriguez fruit bats. Water monitors were tried, but were not 

successful. 

A. capensis: Guenons were housed with A. capensis unsuccessfully at one facility; another facility housed 

them successfully with DeBrazza guenon, with occasional interspecific aggression (R.Willison, personal 


L. canadensis: L. canadensis have been successfully exhibited with beaver. There are unsubstantiated 

reports of exhibiting them with deer, fox, and possibly porcupine in large naturalistic exhibits, but these 

have not been confirmed. Any attempt at mixed species exhibits with the North American river otter 

should take into account their natural inquisitiveness, their semi-aquatic nature, their inclination to climb 

and dig, and their carnivorous diet. 

L. maculicollis: Spotted-necked otters were housed successfully for an extended period with Schmidt’s 

spot-nosed guenon, Allen’s swamp monkey, and François langur. While food was placed in species 

appropriate locations, the otters did eat some of the monkeys’ food. There were some reports of 

intermittent interspecific aggression, generally initiated by the otters in these groupings. An unusual 

aggressive event by a young otter in one of these mixed-exhibits lead to the death of a newborn monkey 

and several days later, the death of the aggressing otter. In this case, the exhibit was re-evaluated and 

discontinued. It should be noted that the otters and monkeys had been exhibited together for several 

years, but the unexpected birth of the guenon infant altered an un-easy truce established between the 

otters and monkeys.



P. brasiliensis: This species should not be kept in mixed-species exhibits due to their highly territorial and 

aggressive nature. There is a record of one unsuccessful attempt at housing giant otter with caiman, 

which was abandoned when a caiman attacked an injured otter and was subsequently removed. 

4.3 Introductions and Reintroductions 

Managed care for and reproduction of animals housed in AZA-accredited institutions are dynamic 

processes. Otters born in or moved between and within institutions require introduction and sometimes 

reintroductions to other animals. It is important that all introductions are conducted in a manner that is 

safe for all otters and humans involved. 

Introductions may take anywhere from a few hours to weeks or months in some cases. There are 

some introductions that will never be successful. In general, these documented, unsuccessful cases 

involved multiple female L. canadensis or P. brasiliensis, male-male introductions in otter species other 

than L. canadensis, and some L. canadensis male-female pairings. However, there is always a risk that 

an introduction will not be successful so all attempts should be monitored closely. 

Otters are very social animals; introductions should be carried out in the standard way, beginning with 

auditory and olfactory introductions, and then moving onto visual and limited tactile contact. Once 

affiliative behavior is observed (e.g., chuckling, rubbing, grunting, “friendly” pawing, and rolling – for 

additional behavior descriptions see Rostain 2000; Reed-Smith 2001; Rostain et al. 2004, Sykes-Gatz 

2005), physical introduction can be attempted, preferably in neutral territory (see exceptions for giant otter 

below). New animals should be allowed to familiarize themselves with their new holding and exhibit prior 

to any physical introductions to other otters/future exhibit-mates. Introducing adult females to other adult 

females is difficult and is generally not recommended for any of the otter species (see Appendix N for 

suggested protocol if female/female introductions are tried). Groups of females that are compatible may 

exhibit aggression towards one another during estrus or be impossible to reintroduce after even a short 

separation (L. canadensis in particular). In cases of aggression during estrus, the females should be 

separated (Schollhamer 1987; J.Reed-Smith, unpublished data). 

Auditory, visual, and olfactory introduction should be successfully completed before attempting 

physical introduction. Successful introductions have been reported as early as one day and have taken as 

long as several months or more. Training animals to station may be beneficial when attempting 

introductions, but there are limited data on its use with otters. 

Reintroductions may need to follow the same steps, particularly if animals have been separated for 

any length of time. When animals are reintroduced to one another, even after short separations, staff 

should carry out each step (auditory, olfactory, visual, limited tactile, then full contact) to ensure animals 

do not experience undue stress or injury. 

L. canadensis: Some introductions will never be successful with L. canadensis, particularly adult female 

to adult female introductions (Reed-Smith 2001) and some adult female to sexually immature male 

introductions. In general, the latter introductions should not be attempted while the female is in estrus; it is 

possible that immature males may be regarded as “female” by adult females leading to aggression on her 

part. Unsuccessful introductions should never progress beyond the screaming, lunging, or fighting stage, 

or result in stressful stand-offs where the animals stay away from one another, scream if the other 

approaches, and may refuse to go on exhibit if the other animal is there. Introductions of breeding pairs 

should be handled as any other, but may move more quickly, particularly during estrus. Introductions of 

all other L. canadensis pairings/groups should be conducted as indicated above. 

Hand-reared L. canadensis may require some time to transfer their focus from humans to otters. The 

easiest way to accomplish this is to have their primary caregiver perform the introductions and stay with 

the animal (free contact situations) or on the other side of the caging (protected contact situations) for the 

initial introductions. Once the hand-reared animal is comfortable with the other otter, their attention often 

transfers immediately to that animal and away from the human caregiver. Hand-reared otters should be 

introduced to other otters as soon after weaning as possible. It is also recommended to raise more than 

one pup together, if possible (Reed-Smith 2004b). Fostering of pups to another nursing female was 

successful following an attempt (Columbus Zoo and Beardsley Zoo). 

L. maculicollis: For male-female introductions, estrus female spotted-necked otters that have previously 

been with a male are easily introduced again to the same male or to a different, experienced male. When 

introducing a sexually mature, but inexperienced male and female, less aggression occurs if the pair is 

introduced when the female is not in estrus (Schollhamer 1987).



P. brasiliensis: P. brasiliensis that are unfamiliar with each other, and those that have been temporarily 

separated (i.e., animals that were previously housed together), should be introduced or reintroduced to 

each other in a gradual, cautious, and closely monitored manner. It is necessary that visual-acousticolfactory 

introductions be conducted before potentially dangerous, full-contact physical introductions are 

attempted. Reports of significant injury and death during improperly conducted giant otter introductions 

have not been uncommon. During introductions/reintroductions, females are more commonly reported to 

have injured/killed males, and even other females, than have males. However, males have been known 

to occasionally injure other males. Females appear to be the more dominant animal during typical 

introductions (this generally holds true even once pairing has been accomplished), and they seem to 

initiate fights more often (Sykes-Gatz 2005). Some giant otters, after 1-2 weeks of visual introduction, 

have been successfully introduced after their first full-contact day; other pairings have taken up to 8 

weeks. 

Initial positive reactions during the first few days of visual and full-contact introductions can be 

misleading. Otters may initially be compatible for some period of time, but show serious aggression later. 

Tolerance, stress, tiredness, aggression, affiliation levels, hunger, etc. should be considered when 

determining timing of initial full-contact sessions, when to increase contact time, and when to decrease or 

stop contact. Some tension and minor, non-harmful fights should be allowed, but temporary separation is 

required before serious fighting develops. During introductions, it is advisable to use fence barriers with 

~2cm x 2cm (1" x 1") mesh size or cover existing fences with material of this mesh size. This will prevent 

fence fighting and/or injury to body parts. It is advisable to conduct introductions in areas of sufficient size, 

allowing for proper interaction between the otters; Sykes-Gatz (2005) recommends an introduction fence 

of at least 5m (16.4ft) in length. During introductions, the otters should be provided with sufficient living 

space to allow for their privacy and isolation if desired. The International Giant Otter Studbook Husbandry 

and Management Information and Guidelines (Sykes-Gatz 2005) provides greater detail on the 

recommended introduction procedure for this species.

5.1 Nutritional Requirements 

A formal nutrition program is recommended to meet the 

behavioral and nutritional needs of all animals (2.6.2). Diets 

should be developed using the recommendations of veterinarians 

as well as AZA Taxon Advisory Groups, Species Survival 

Plans®, and Nutrition Advisory Groups 

(www.nagonline.net/feeding_guidelines.htm). Diet formulation 



Chapter 5. Nutrition 


(2.6.2) A formal nutrition program is 

recommended to meet the behavioral and 

nutritional needs of all species and 

specimens within the collection. 

criteria should address the otters’ nutritional needs, feeding ecology, as well as individual and natural 

histories to ensure that species-specific feeding patterns and behaviors are stimulated. 

The target nutritional values for otters are based on several sources. The cat is typically used to 

establish nutrient guidelines for carnivorous animals. The National Research Council (NRC) (1986, 2006), 

Association of American Feed Control Officials (AAFCO) (1994), and Waltham Center for Pet Nutrition 

(Earle & Smith 1993) have provided recommendations for cats. A limited amount of information is 

provided by the NRC publication for mink and foxes (1982), which represents the requirements of another 

mustelid species. The target nutrient values presented (Maslanka & Crissey 1998) are a range of values 

reported from various references. As new information becomes available, these ranges will change to 

reflect knowledge gained. Table 3 lists dietary nutrient ranges for otters. 

Table 4 contains updated information on feline nutritional requirements based on NRC 

recommendations published in 2006. The original target values have been retained for comparison. See 

Dierenfeld et al. (2002) for information on nutrient composition of whole vertebrate prey. Appendix H 

provides a description of the nutrients listed in these tables. 

Diet formulation should account for animal preferences, body weight, exercise, physical condition, 

environmental/seasonal changes, behavioral considerations, diet item availability, gastrointestinal tract 

morphology, and actual nutrient requirements. 

Primarily piscivorous, otters have high metabolic rates, rapid digestion, and have been found to 

spend 41-60% of their time involved in feeding or foraging activities (Hoover & Tyler 1986; Davis et. al 

1992; Kruuk 1995; J.Reed-Smith, unpublished data). Duplaix-Hall (1975) found that otters (unidentified 

species) in the wild rarely ate more than about 500g of food at a time and that they consumed 

approximately 20% of their own body weight daily. Kruuk (1995) reviewed his and other study results 

indicating that ex-situ populations of Lutra lutra consuming between 11.9-15% of their body weight 

maintained a healthy weight. Ben-David et al. (2000, 2001a and b) reported success using 10% of a L. 

canadensis’ (ex-situ population) body weight as a guide for the basis of their maintenance diet. See 

section 5.2 for sample diets for the various otter species.

Table 3: Target dietary nutrient ranges for otters (dry matter basis). 

Item Target nutrient range* 

Energy, kcal/g 3.6-4.0 

Crude Protein, % 24-32.5 

Fat, % 15-30** 

Vitamin A, IU/g 3.3-10*** 

Vitamin D. IU/g 0.5-1.0 

Vitamin E, mg/kg 30-120 a 

Thiamin, mg/kg 1-5 a 

Riboflavin, mg/kg 3.7-4.0 

Pantothenic Acid, mg/kg 5-7.4 

Niacin mg/kg 9.6-40 

Pyridoxine, mg/kg 1.8-4.0 

Folic acid, mg/kg 0.2-1.3 

Biotin, mg/kg 0.07-0.08 

Vitamin B-12, mg/kg 0.02-0.025 

Choline, mg/kg 1000-3000 

Calcium, % 0.6-0.8 b 

Phosphorus % 0.6 b 

Potassium, % 0.2-0.4 

Sodium, % 0.04-0.06 

Magnesium, % 0.04-0.07 

Zinc, % 50-94 

Copper, mg/kg 5-6.25 

Manganese, mg/kg 5.0-9.0 

Iron, mg/kg 80-114 

Iodine, mg/kg 1.4-4.0 



* Target nutrient ranges expressed on a dry matter basis derived from requirements for domestic cats (NCR 1986), AAFCO 

recommendations (1994), Waltham Center for Pet Nutrition recommendations (Earle & Smith 1993), and requirements for mink and 

foxes (NCR 1982). 

** The fat content of fish commercially available in N.A. typically ranges from 5–40% (Maslanka & Crissey 1998), and N.A. river 

otters have been maintained on diets containing 24-30% fat (Reed-Smith 1994), thus an appropriate range for fat appears to fall 

between 15-30%. 

*** The vitamin A requirement for cats is 10 IU/g (dry matter basis; NRC 1986), which represents the upper bound of the range. 

However, free-ranging N.A. otters consume a higher proportion of fish and may have a higher tolerance for vitamin A due to the high 

levels, which occur in their natural diet. 

a When mostly fish diets are offered, the presence of unsaturated fatty acids and thiaminases causes the breakdown of these 

vitamins. Thus, dietary levels of 400 IU vitamin E/kg of dry diet and 100-120mg thiamin/kg of dry diet or 25-30mg thiamine/kg fresh 

weight as fed basis are recommended (Engelhardt & Geraci 1978; Bernard & Allen 1997). 

b The recommended Ca:P ration is between 1:1 and 2:1

Table 4: Target nutrient ranges for carnivorous species (dry matter basis) 

Nutrient 

NRC 1986 

Cat 1 

Maintenance 

Growth 

NRC 2006 Cat 2 Arctic 

fox 3 

Maintenance 

Gestation 

Lactation 

Maintenance 



Mink 4 Carniv 5 

Protein (%) 24-30 22.5 20 21.3-30 19.7-29.6 21.8-26 19.7-30 

Fat (%) 9.0-10.5 9.0 9.0 15.0 -- -- 9-15 

Linoleic Acid (mg/kg) 0.5 0.55 0.55 0.55 -- -- 0.5-0.55 

Vitamin A (IU/g) 3.3-9.0 3.55 3.55 7.5 2.44 5.9 2.44-9 

Vitamin D (IU/g) 0.5-0.75 0.25 0.25 0.25 -- -- 0.25-0.75 

Vitamin E (mg/kg) 27-30 38.0 38.0 38.0 -- 27.0 27-38 

Vitamin K (mg/kg) 0.1 1.0 1.0 1.0 -- -- 0.1-1 

Thiamin (mg/kg) 5.0 5.5 5.6 5.5 1.0 1.3 1-5.6 

Riboflavin (mg/kg) 3.9-4.0 4.25 4.25 4.25 3.7 1.6 1.6-4.25 

Niacin (mg/kg) 40-60 42.5 42.5 42.5 9.6 20.0 9.6-60 

Pyridoxine (mg/kg) 4.0 2.5 2.5 2.5 1.8 1.6 1.6-4 

Folacin (mg/kg) 0.79-0.8 0.75 0.75 0.75 0.2 0.5 0.2-0.8 

Biotin (mg/kg) 0.07-0.08 0.075 0.075 0.075 -- 0.12 0.07-0.12 

Vitamin B12 (mg/kg) 0.02 0.022 0.022 0.022 -- 0.032 0.02-0.032 

Pantothenic acid (mg/kg) 5.0 6.25 6.25 6.25 7.4 8.0 5-8 

Choline (mg/kg) 2400 2550 2550 2550 -- -- 2400-2550 

Calcium (%) 0.8-1.0 0.8 0.29 1.08 0.6 0.3-0.4 0.29-1.08 

Phosphorus (%) 0.6-0.8 0.72 0.26 0.76 0.6 0.3-0.4 0.26-0.8 

Magnesium (%) 0.03-0.08 0.04 0.04 0.06 -- -- 0.03-0.08 

Potassium (%) 0.4-0.6 0.4 0.52 0.52 -- -- 0.4-0.6 

Sodium (%) 0.05-0.2 0.14 0.068 0.132 -- -- 0.05-0.2 

Iron (mg/kg) 80.0 80.0 80.0 80.0 -- -- 80 

Zinc (mg/kg) 50-75 75.0 75.0 60.0 -- -- 50-75 

Copper (mg/kg) 5.0 8.4 5.0 8.8 -- -- 5-8.8 

Manganese (mg/kg) 5.0 4.8 4.8 7.2 -- -- 4.8-7.2 

Iodine (mg/kg) 0.35-0.42 2.2 2.2 2.2 -- -- 0.35-2.2 

Selenium (mg/kg) 0.1 0.4 0.4 0.4 -- -- 0.1-0.4 

1 

NRC (1986), Legrand-Defretin and Munday (1993), AAFCO (1994). All numbers are based on requirement set for maintenance. 

2 

Dog and Cat NRC (2006). 

3 

NRC (1982). Protein is range of growth and maintenance; vitamins are for growth, and minerals for growth and maintenance. 

4 

NRC (1982). Protein is for maintenance, vitamins are for weaning to 13 weeks and minerals are a range of growing and 

maintenance. 

5 

Combination of cat, mink, and fox 

Changing Nutrient Requirements – Age: An animal’s diet should be developed to maintain optimal 

weight or weight gain and normal physical development for a young animal. Diets for young or senescent 

adults should take into account their activity level, dental development and/or condition. 

P. brasiliensis: In an ex-situ population study, Carter and Rosas (1997) determined that an adult 

consumed roughly 10% (range 6-16%) of their body weight daily and a sub-adult consumed 13.4% (range 

8-18.9%). Earlier studies (Zeller 1960; Best 1985) reported similar findings with adults and sub-adults 

daily consuming 7-9.6% and 12.9% of their body weight, respectively. Amounts eaten can vary with air 

temperature and activity level changes, but if food is refused for one day, this could be a sign of sickness. 

Excess weight gain or loss and daily amounts and food types eaten should be monitored and recorded 

(Sykes-Gatz 2005). 

Changing Nutrient Requirements – Reproduction: There is an increased need for energy during 

lactation. Tumanov & Sorina (1997) supported the use of high-energy diets for lactating female mustelids. 

Fat is the most concentrated source of energy in the diet. For lactating females, fat levels in the diet may 

be increased to support lactation (see below for exceptions) and also to provide increased energy to 

minimize mobilization of body stores and metabolic stress associated with milk production. Diet increases 

Maintenance 

All



for lactating otters should be based on past experiences with individual otters and/or observed body 

weight loss (mobilization of tissue to support lactation). To date, institutions have typically increased the 

amount of fish offered a lactating female versus simply increasing the fat content by switching the type of 

food offered. An increase of 10-30% is the accepted rule. 

P. brasiliensis: Hagenbeck and Wünnemann (1992) reported that lactating females at the Hagenbeck 

Tierpark generally increased their food consumption from 4.41-6.61 lbs/day to 13.23lbs/day (2-3kg/day to 

6kg/day). They also reported increasing vitamin supplements during pregnancy/lactation and calcium 

supplementation during lactation (Sykes-Gatz 2005). 

The energy requirements of a pair of otters, including a pregnant female, at the Philadelphia Zoo also 

increased during pregnancy and lactation. At this time, the energy intake of the pair increased to 

246kcal/kg BW 0.75 (~2.75kg fish/animal fed at a ratio of 1:2 low- to high-fat fish). Fifty days postpartum 

and with one surviving pup, the intake of the pair was 236kcal/kg BW 0.75 (~3kg fish/animal fed at a ratio of 

1:4 low- to high-fat fish). The female exhibited a preference for herring, trout, and catfish (K.Lengel, 

personal communication). It appears that feeding behaviors of ex-situ populations of reproductive P. 

brasiliensis mimic those of their wild counterparts. Rosas et al. (1999) found that during the birthing 

season, the diet of wild otters included a higher proportion of fish in the order Siluriformes (catfishes), 

which are higher in fat (37-41% fat DMB – Silva 1993) than fish in the order Percoidei (perch) (22-31% fat 

DMB – Twibell & Brown 2000),which are commonly fed on in the wild. Siluriformes are also higher in fat 

than Chiclidae (tilapia) (21-32% fat DMB – Toddes 2005-2006 analysis), which are the low-fat fish 

commonly fed to otters at the Philadelphia Zoo. 

Seasonal Changes in Nutritional Needs: An animal’s weight should be monitored regularly and diets 

adjusted accordingly. Some institutions report seasonal changes in appetite of some otters, but not in the 

majority of animals. Further research in this area is required. An animal’s weight should be regularly 

monitored and diets adjusted accordingly. At this time, further research into seasonal nutritional 

requirements is required. 

P. brasiliensis: The energy needs of P. brasiliensis are very dependent on their life stage, social grouping, 

and the ambient temperature of the environment. At the Philadelphia Zoo, an average energy intake of 

173kcal/kg BW 0.75 (~2kg of fish fed at a ratio of 3:1 low- to high-fat fish) was adequate to maintain a single 

adult otter within a target weight range during the warmer months of the year (K.Lengel, personal 

experience). 

When maintained with a mate, the same animal required an increased energy intake from 173kcal/kg 

to 201kcal/kg BW 0.75 (2.75kg of fish fed at a ratio of 3:1 low- to high-fat fish) during warmer months and 

went as high as 243kcal/kg BW 0.75 (2.75kg of fish fed at a ratio of 2:1 low- to high-fat fish) during cooler 

months (K.Lengel, personal experience). 

A group of two adults, an 18-month old juvenile male, and three 6-month-old pups were successfully 

maintained during the summer season on an average energy allotment for the group of 545kcal/kg BW 0.75 

(~6kg fish/animal fed at a ratio of 1.25:1 low- to high-fat fish). This energy allotment exceeded that of 

previous intake studies by almost double. However, the group was extremely active and primarily 

comprised of growing adolescent animals. 

Weight Loss: While otters should carry some body fat and not be kept artificially thin, they are prone to 

gaining excessive weight in zoos and aquariums. Tarasoff (1974) reported subcutaneous fat deposits 

primarily at the base of the tail and caudally on the rear legs, with smaller deposits around the genitalia 

and in the axillary regions. There are several ways to approach formulating a weight loss diet for otters. 

Depending on the food items available, the feeding situation (fed alone or in a group), and the amount of 

weight loss desired, one or more of the following approaches may be appropriate. 

Feed less total food: By reducing the amount of total food offered, weight loss may occur. This practice is 

confounded by the aggression observed in most otters, and particularly A. cinereus and P. brasiliensis 

groups, around feeding time and the potential for this to increase when less food is offered. 

Add more water to the diet: By providing a diet that contains more moisture, the total calories in the diet 

are diluted and this may allow for weight loss. The otter can consume the same amount of total diet, but 

will actually be consuming fewer calories.



Increase the “bulk” of the diet: By adding indigestible or lower calorie items to the diet, the total “bulk” of 

the diet can be increased, effectively diluting the calories in the diet. The otter can consume the same 

amount of total diet, but will actually be consuming fewer calories. 

Offer lower calorie items: Lower calorie items can be substituted in the diet. For example, fish varies in 

energy content from species to species. If weight loss is desired, a leaner fish, such as Pollock, could be 

substituted for a fattier fish, such as herring or capelin, to reduce total calories in the diet. This would be 

the preferred method for all otter species fed fish. 

5.2 Diets 

The formulation, preparation, and delivery of all diets must 

be of a quality and quantity suitable to meet the animal’s 

psychological and behavioral needs (2.6.3). Food should be 

purchased from reliable, sustainable, and well-managed 

sources. The nutritional analysis of the food should be regularly 

tested and recorded. 

Food preparation must be performed in accordance with all 

relevant federal, state, or local regulations (2.6.1). Meat 

processed on site must be processed following all USDA 

standards. 

If browse plants are used within the animal’s diet or for 

enrichment, all plants must be identified and assessed for 

safety. The responsibility for approval of plants and oversight of 

the program should be assigned to at least one qualified 

individual (2.6.4). The program should identify if the plants have 

been treated with any chemicals or near any point sources of 

pollution and if the plants are safe for the species. If otters have 

access to plants in and around their exhibits, there should be a 

staff member responsible for ensuring that toxic plants are not 

available. Before any plants are placed in or near otter exhibits, 

they should be vetted through relevant management staff (e.g. 


(2.6.3) Animal diets must be of a quality 

and quantity suitable for each animal’s 

nutritional and psychological needs. Diet 

formulations and records of analysis of 

appropriate feed items should be 

maintained and may be examined by the 

Visiting Committee. Animal food, 

especially seafood products, should be 

purchased from reliable sources that are 

sustainable and/or well managed. 


(2.6.1) Animal food preparations must 

meet all local, state/provincial, and federal 

regulations. 


(2.6.4) The institution should assign at 

least one person to oversee appropriate 

browse material for the collection. 

curator, veterinarian, grounds keeper, county poison control officer, etc.). Plants considered toxic to 

humans or other animals should be considered toxic to otters. Loquat (Weber and Garner 2002) 

consumption has proven fatal to Asian small-clawed otters (see useful Veterinary References). Otters are 

obligate carnivores but they will eat some vegetative matter such as berries and/or consume vegetation 

or other foreign material out of boredom or while exploring their environment. 

Sample Diets: The one best diet for any of the otters of ex-situ populations has not been found and 

requires further research. However, current recommendations for all but A. cinereus are that a variety of 

fish species should be offered 3-4 times a week, preferably daily. Currently the AZA Otter SSP 

recommends a specific diet for A. cinereus (see below) and that P. brasiliensis should be offered fish 

daily as their main diet. The AZA Small Carnivore TAG chair should be contacted for specifics regarding 

the A. cinereus diet. 

A. cinereus: The following food items represent the recommended daily diet per animal for A. cinereus 

(AZA Otter SSP recommendation 2006). The items are given as percentage of diet fed: 

• 54.5% commercial canned diet designed to meet the nutrient requirements for domestic cats and 

control occurrence of calcium oxalate uroliths (e.g., Hill’s x/d ® , IAMS Moderate pH/O ® ). 

• 2.5% commercial dry food to meet nutrient requirements for domestic cats and control occurrence 

of calcium oxalate uroliths (e.g., Hill’s x/s ® , IAMS Moderate pH/O ® ). 

• 17.4% capelin 

• 24.6% lake smelt 

• 1% cricket and meal worms 

• 100 IU vitamin E per kg of fish offered 

• 25-35mg thiamin per kg of fish offered 

A. capensis: The following food items represent a sample daily diet per animal for A. capensis in ex-situ 

environments.

• 182g Dallas Crown Carnivore Diet 

• 908g fish (herring, capelin, sardines, two choices daily) 

• 1.5 dozen large crayfish a week 

• 30g lamb & rice dry dog food 

• 90 IU vitamin E 

• 6.25mg thiamin 



L. canadensis: The amounts of food items in the sample diet below are based on achieving a target 

weight for otters. The diet should be fed at least three times a day and 4-5 times if possible. These 

additional feedings can consist of the fish, rib bones, and enrichment/training feeds. 

• 155g commercially prepared feline diet, 2 x day, 7 days a week 

• 112g capelin, 1 x day 

• 120g smelt, 1 x day 

• 135g trout, 1 x day 

• ½ medium carrot, 1 x day scattered 

• 2 rib bones, ox tail, or similar 3 x week 

• 25-35mg thiamin per kg of fish offered 

• 100 IU vitamin E per kg of fish 

Only good quality, mostly fresh water fish, low in thiaminase and fat should be offered (Wünnemann 

1995a). The fish source(s) and/or vendor(s) should be examined closely to assess their handling 

practices, ensure that HACCP guidelines are being met, and that the fish is considered human grade. 

Historical use of a type of fish by zoos and aquariums does not ensure it is an adequate diet ingredient, 

and only careful inspection of handling practices and the fish itself ensures consistent safety and quality. 

Most diets currently include horsemeat products, or alternative beef-based products which are available 

in addition to nutritionally complete dry and wet cat foods. The following sample diet is recommended for 

L. canadensis: 

• 13.5% capelin 

• 14.5% smelt 

• 16.3% herring 

• 18.2% carrots 

• 37.5% nutritionally complete cat food or beef-based product (IAMS ® cat food used for analysis) 

• 2 bones, 3 per week (rib, ox/horse tail, or similar) 

• 25-35mg thiamin and 100 IU vitamin E per kg of fish fed 

L. maculicollis: The following food items represent a sample daily diet per animal for L. maculicollis. The 

animals should be fed at least twice per day: 

• 50g Iams ® Less Active Cat Kibble 

• 150g Natural Balance zoo carnivore diet 

• 150g trout (3 x week) 

• 120g squid (3 x week) 

• Yams and carrots offered in small amounts. 

P. brasiliensis: A variety of good quality, fresh-water fish low in thiaminase and fat should be offered as 

the main diet (Wünnemann 1995a). Saltwater fish, high in fat, should only be offered occasionally. This 

species should be fed 3-5 times daily. Typically, 2-3kg (4.4-6.6lbs) of fish should be fed daily to each 

adult. Results of a survey of facilities housing this species indicate that all of these institutions offer fish 

daily (thawed, frozen, live, and/or freshly caught) as the main diet. Fish species offered include the 

following: rainbow trout (Salmo gairdneri), carp (cyprinus carpio), river fish (unidentified), tilapia, redeye 

(Rutilus rutilus), common bream (Abramis brama), herring* (Culpea harengus), mackerel* (Scomber 

scombrus), felchen (Coregonus albula), and channel catfish (Ictalurus punctatus). Fish species marked 

with an asterisk (*) can be used as a training reward or for vitamin delivery. If thawed frozen fish 

constitute the bulk of the diet the otters should be given supplementary B1 (thiamine) and vitamin E. 

Supplements should be fed separately from the main feedings by at least 2 hours (Sykes-Gatz 2005). 

For all otters: Fish types containing high thiaminase and/or high polyunsaturated fat levels should be 

avoided as they can cause malnutrition, sickness, and even death (Merck 1986). Diets containing fish



high in thiaminase can lead to thiamin (vitamin B1) deficiency in the otters fed this diet (Merck 1986). The 

process of fish storage (freezing), thawing, and preparation, can lead to fish nutrient loss, particularly 

vitamins B1 and E, and especially in fish with high fat and/or high thiaminase content (Crissey 1998; 

Merck 1986). Vitamin supplements, especially vitamin B1 (thiamin), vitamin E, and a multivitamin, should 

be added when fish is the main diet. The recommended vitamin supplementation regime for fish eating 

animals is as follows: 

• Thiamin: 25-30mg/kg fish fed, fresh weight as fed basis (Bernard & Allen 1997) 

• Vitamin E: 400 IU/kg dry weight basis (Engelhardt & Geraci 1978) 

Based on the information above, the following food items represent a sample diet for giant otters 

(Sykes-Gatz 2005): 

• 2-3kg (4.4-6.6lbs) fish/day/adult 

• 400 IU vitamin E daily 

• 100mg vitamin B1 daily 

• Multi-vitamin/mineral supplement 3 x week 

Feeding Schedule: Due to their naturally nutrient dense diet, reliance on fat as a source of energy, rapid 

transit time of food through the intestinal tract, feeding style of frequent, small amounts, and generally 

high activity level – it is recommended that otters be fed at least twice a day and preferably three or more 

times daily (including enrichment or training feeds). P. brasiliensis should be fed 3-5 times per day. 

Frequent feeding prevents consumption of spoiled food, accommodates their rapid digestion (Ormseth & 

Ben-David 2000), and can stimulate increased activity in these generally active and curious species. 

In addition to feeding smaller amounts frequently, it is recommended that a portion of the daily diet be 

fed as part of enrichment or husbandry training activities. At least one of the daily feedings, or part of a 

feeding, should be scattered to encourage foraging (except for giant otter). Timing of foraging 

opportunities and items offered should be varied to prevent habituation. All uneaten food should be 

removed before it spoils; this may be daily or more frequent in warm climates or seasons. 

P. brasiliensis: Food for P. brasiliensis should not be scatter fed, as they do not forage on land and nonliving 

food left uneaten in pools can be difficult to find. A portion of the daily diet can be used for daily 

training sessions with this species. 

Food Variability: Otters should routinely be offered a variety of fish either as part of their diet or as 

enrichment. Reliance on multiple fish species, versus one or two, will prevent animals from developing 

strong preferences and help in switching them over to new sources if one fish type becomes unavailable. 

With the exception of P. brasiliensis (see below), otters will sample a variety of food groups, 

especially if introduced to them at an early age; cat kibble, worms, crickets, vegetables, berries, mice, 

chicks, etc., can all be added to the diet as enrichment. Due to the possible formation of uroliths, foods 

high in calcium oxalates should be avoided (e.g., beans, celery, leafy greens, sweet potato, berries, 

peanuts, among others), particularly for A. cinereus. The use of these items for enrichment scatter feeds 

for North American river otter is acceptable on a limited basis, but the overall nutrient and caloric intake, 

body weight of the animal(s), and condition of the animal(s) should be taken into consideration. 

All otters will benefit from receiving live fish/crayfish (from approved sources), at least as enrichment 

on a weekly basis. Whole fish should be the sole dietary item offered to P. brasiliensis and should 

comprise a portion of the daily diet of all other species. 

A. cinereus: The AZA Otter SSP has specific diet recommendations for this species that should be 

obtained from the AZA Otter SSP Chair 

A. capensis, L. canadensis, and L. maculicollis: It is recommended that fish constitute at least a portion of 

the daily diet offered these species. Hard dietary items should be routinely incorporated for dental health. 

These can include: hard kibble, crayfish, crabs, chicken necks, ox/horse tails, partially frozen fish, bony 

fish, day-old chicks, mice, rib bones, canine dental bones, or similar items. 

P. brasiliensis: Staib (2002) reports that wild giant otters almost exclusively eat fish. In the wild, fish from 

the suborders Characoidei (characins), Percoidei (perch), and Siluroidei (catfish) make up the majority of 

the giant otter diet (Carter & Rosas 1997). A variety of good quality, fresh-water fish low in thiaminase 

and fat should be offered as their main diet (Wünnemann 1995a). Saltwater fish, high in fat, should only 

be offered occasionally. Gravid fish have caused diarrhea and appetite loss, and fish eggs should be



removed before feeding (V.Gatz, personal observation). This species should be fed 3-5 times daily 

(typically 2-3kg [4.4-6.6lbs] fish/day/adult) (Sykes-Gatz 2005). A small amount of left over food is common 

and desirable to ensure all members of the group receive their portion and to avoid fights over fish. 

Uneaten fish should be regularly removed to prevent the otters from consuming spoiled food (Sykes-Gatz 

2005). The strategy of feeding animals multiple times per day and using at least some feedings as 

training sessions has been successful at maintaining animal weights and maintaining low levels of food 

aggression in the group (Toddes 2005-2006). 

Species-appropriate Foraging and Feeding: Live fish and crustaceans can and should be provided, if 

possible, on a regular basis. However, due to the risks of live fish or crayfish transmitting disease or 

parasites, policies regarding the feeding of live prey should be established by each facility. If these items 

are used, they should be obtained only from known, institutionally approved sources. Where live prey are 

used, provisions in the exhibit should be made to allow these prey species a place to hide from the otters, 

thus forcing the otters to use their hunting skills and extending the time of activity. 

There also are a variety of puzzles and other feeding devices described in the literature that can be 

adapted for use in river otters. Alternatively, feeding tubes can be built into exhibits that randomly release 

live prey or food items into the exhibit. See Chapter 8, section 8.2 for other enrichment items used, 

including non-food items. 

5.3 Nutritional Evaluations 

The Otter SSP is currently beginning work on a body-condition matrix that can be used to help assess 

proper weight and condition for otters (See Appendix O for working matrix draft). At this time there are no 

known tools for performing clinical nutritional evaluations of otters; this would be a useful area for future 

research.

6.1 Veterinary Services 

AZA General Policies: Veterinary services are a vital 

component of excellent animal care practices. A full-time staff 

veterinarian is recommended, however, in cases where this is 

not practical, a consulting/part-time veterinarian must be under 

contract to make at least twice monthly inspections of the animal 

collection and to any emergencies (2.1.1). Veterinary coverage 

must also be available at all times so that any indications of 

disease, injury, or stress may be responded to in a timely 

manner (2.1.2). All AZA-accredited institutions should adopt the 

guidelines for medical programs developed by the American 

Association of Zoo Veterinarians (AAZV) 

(www.aazv.org/associations/6442files/zoo_aquarium_vet_med_ 

guidelines.pdf). The current Veterinary Advisor for the AZA Otter 

SSP is Dr. Gwen Myers. 

Protocols for the use and security of drugs used for 

veterinary purposes must be formally written and available to 

animal care staff (2.2.1). Procedures should include, but are not 

limited to: a list of persons authorized to administer animal 

drugs, situations in which they are to be utilized, location of 

animal drugs and those persons with access to them, and 

emergency procedures in the event of accidental human 

exposure. 

Animal recordkeeping is an important element of animal care 

and ensures that information about individual otters and their 

treatment is always available. A designated staff member should 

be responsible for maintaining an animal record keeping system 

and for conveying relevant laws and regulations to the animal 

care staff (1.4.6). Recordkeeping must be accurate and 

documented on a daily basis (1.4.7). Complete and up-to-date 

animal records must be duplicated and retained in a fireproof 

container within the institution (1.4.5) as well as be duplicated 

and stored at a separate location (1.4.4). Giant otter are a 

USFWS and IUCN listed endangered species, all relevant record 

keeping regulations should be followed. 

6.2 Identification Methods 

AZA Policies: Ensuring that animals are identifiable through 

various means increases the ability to care for individuals more 

effectively. Animals must be identifiable and have corresponding 

ID numbers whenever practical, or a means for accurately 

maintaining animal records must be identified if individual 

identifications are not practical (1.4.3). 

AZA Otter SSP Identification Recommendations: The AZA 

Otter SSP recommends that all otters be identified as soon as 

possible after birth with a transponder chip. Chips have been 

placed subcutaneously over the bridge of the nose/forehead 

area, SQ/IM in the intrascapular area at the base of the ears, 

and many institutions have placed them between the scapulas 

(recommended placement for giant otter). Placement in all of 

these areas has been met with success and failure (migration, 

loss, unable to read them as planned). At this time, the AZA 

Otter SSP recommends the forehead area as the preferred area 



Chapter 6. Veterinary Care 


(2.1.1) A full-time staff veterinarian is 

recommended. However, the Commission 

realizes that in some cases such is not 

practical. In those cases, a 

consulting/part-time veterinarian must be 

under contract to make at least twice 

monthly inspections of the animal 

collection and respond as soon as 

possible to any emergencies. The 

Commission also recognizes that certain 

collections, because of their size and/or 

nature, may require different 

considerations in veterinary care. 


(2.1.2) So that indications of disease, 

injury, or stress may be dealt with 

promptly, veterinary coverage must be 

available to the animal collection 24 hours 

a day, 7 days a week. 


(2.2.1) Written, formal procedures must 

be available to the animal care staff for 

the use of animal drugs for veterinary 

purposes and appropriate security of the 

drugs must be provided. 


(1.4.6) A staff member must be 

designated as being responsible for the 

institution's animal record-keeping 

system. That person must be charged 

with establishing and maintaining the 

institution's animal records, as well as 

with keeping all animal care staff 

members apprised of relevant laws and 

regulations regarding the institution's 

animal collection. 


(1.4.7) Animal records must be kept 

current, and data must be logged daily. 


(1.4.5) At least one set of the institution’s 

historical animal records must be stored 

and protected. Those records should 

include permits, titles, declaration forms, 

and other pertinent information. 


(1.4.4) Animal records, whether in 

electronic or paper form, including health 

records, must be duplicated and stored in 

a separate location.

of placement for all but the giant otter (see below); this location 

should make the chip easy to read when the animal comes to 

the front of the cage. The intrascapular area should be used as 

an alternative (this is the most frequently used location reported 

by member institutions). However, transponders placed in the 

intrascapular area can migrate and may be broken or lost during 

fighting and breeding attempts. Placement location of the 

transponder chip should be recorded in the animal’s medical 

record. 



P. brasiliensis: For this species it is recommended that 

transponder chips be placed in the neck behind the left ear or the intrascapular area. Transponders 

should be placed deeply under the skin or intramuscularly to decrease the risk of lost or damaged 

transponders (C.Osmann, personal communication). 

AZA Acquisition/Disposition Policies: AZA member 

institutions must inventory their population at least annually and 

document all animal acquisitions and dispositions (1.4.1). 

Transaction forms help document that potential recipients or 

providers of the otters should adhere to the AZA Code of 

Professional Ethics, the AZA Acquisition/Disposition Policy (see 

Appendix B), and all relevant AZA and member policies, 

procedures and guidelines. In addition, transaction forms must 

insist on compliance with the applicable laws and regulations of 

local, state, federal and international authorities. All AZAaccredited 

institutions must abide by the AZA Acquisition and 

Disposition policy (Appendix B) and the long-term welfare of 

animals should be considered in all acquisition and disposition 

decisions. All species owned by an AZA institution must be 

listed on the inventory, including those animals on loan to and 

from the institution (1.4.2). Standard institutional and AZA approved acquisition and disposition policies 

and forms are used for otters. The AZA Otter SSP and relevant species coordinator should be notified 

when otters are acquired or transferred between institutions. 

6.3 Transfer Examination and Diagnostic Testing Recommendations 

AZA Policies: The transfer of animals between AZA-accredited institutions or certified related facilities 

due to SSP or PMP recommendations occurs often as part of as concerted effort to preserve these 

species. These transfers should be done as altruistically as 

possible and the costs associated with specific examination and 

diagnostic testing for determining the health of these otters 

should be considered. 

AZA Otter SSP Recommendations: See Section 6.4 

Quarantine. 

6.4 Quarantine 

AZA Policies: AZA institutions must have holding facilities or 

procedures for the quarantine of newly arrived animals and 

isolation facilities or procedures for the treatment of sick/injured 

animals (2.7.1). All quarantine, hospital, and isolation areas 

should be in compliance with AZA standards/guidelines (2.7.3; 

Appendix C). All quarantine procedures should be supervised by 

a veterinarian, formally written and available to staff working with 

quarantined animals (2.7.2). If a specific quarantine facility is not 

present, then newly acquired otters should be kept separate from 

the established collection to prohibit physical contact, prevent 

disease transmission, and avoid aerosol and drainage 


(1.4.3) Animals must be identifiable, 

whenever practical, and have 

corresponding ID numbers. For animals 

maintained in colonies or other animals 

not considered readily identifiable, the 

institution must provide a statement 

explaining how record keeping is 

maintained. 


(1.4.1) An animal inventory must be 

compiled at least once a year and include 

data regarding acquisitions and 

dispositions in the animal collection. 


(1.4.2) All species owned by the 

institution must be listed on the inventory, 

including those animals on loan to and 

from the institution. In both cases, 

notations should be made on the 

inventory. 


(2.7.1) The institution must have holding 

facilities or procedures for the quarantine 

of newly arrived animals and isolation 

facilities or procedures for the treatment 

of sick/injured animals. 


(2.7.3) Quarantine, hospital, and isolation 

areas should be in compliance with 

standards or guidelines adopted by the 

AZA. 


(2.7.2) Written, formal procedures for 

quarantine must be available and familiar 

to all staff working with quarantined 

animals. 

contamination. If the receiving institution lacks appropriate facilities for quarantine, pre-shipment



quarantine at an AZA or AALAS accredited institution may be applicable. Local, state, or federal 

regulations that are more stringent than AZA Standards and recommendation have precedence. 

AZA institutions must have zoonotic disease prevention 

procedures and training protocols established to minimize the 

risk of transferable diseases (11.1.2) with all animals, including 

those newly acquired in quarantine. Keepers should be 

designated to care only for quarantined animals if possible. If 

keepers have to care for both quarantined and resident otters of the same class, they should care for the 

quarantined animals only after caring for the resident animals. Equipment used to feed, care for, and 

enrich otters in quarantine should be used only with these otters. If this is not possible, then all items 

should be appropriately disinfected, as designated by the veterinarian supervising quarantine before use 

with resident otters. 

Quarantine durations span of a minimum of 30 days (unless otherwise directed by the staff 

veterinarian). If additional carnivores are introduced into their corresponding quarantine areas, the 

minimum quarantine period should begin over again. However, the addition of mammals of a different 

order will not require the re-initiation of the quarantine period. 

During the quarantine period, specific diagnostic tests should be conducted with each otter if possible 

or (see Appendix C). A complete physical, including a dental examination if applicable, should be 

performed. Otters should be evaluated for ectoparasites and treated accordingly. Blood should be 

collected, analyzed and the sera banked in either a -70ºC freezer or a frost-free -20ºC freezer for 

retrospective evaluation. Fecal samples should be collected and analyzed for gastrointestinal parasites 

and the otters should be treated accordingly. Vaccinations should be updated as appropriate, and if the 

vaccination history is not known, the otter should be treated as immunologically naive and given the 

appropriate series of vaccinations. 

A tuberculin testing and surveillance program must be 

established for animal care staff as appropriate to protect both the 

health of both staff and animals (11.1.3). Depending on the 

disease and history of the otters, testing protocols for animals 

may vary from an initial quarantine test to yearly repetitions of 

diagnostic tests as determined by the veterinarian. TB testing is 

not routinely performed on otters. Animals should be permanently 

identified by their natural markings or, if necessary, marked when anesthetized or restrained (e.g., tattoo, 

ear notch, ear tag, etc.). Release from quarantine should be contingent upon normal results from 

diagnostic testing and two negative fecal tests that are spaced a minimum of two weeks apart. Medical 

records for each animal should be accurately maintained and easily available during the quarantine 

period. 

Although every living animal dies at some point, otters which 

die during the quarantine period should have a necropsy 

performed to determine the cause of death and the subsequent 

disposal of the body must be done in accordance with any local or 

federal laws (2.5.1). Necropsies should include a detailed external 

and internal gross morphological examination and representative 

tissue samples form the body organs should be submitted for 

histopathological examination. 


(11.1.2) Training and procedures must be 

in place regarding zoonotic diseases. 


(11.1.3) A tuberculin testing and 

surveillance program must be 

established for appropriate staff in order 

to ensure the health of both the 

employees and the animal collection. 


(2.5.1) Deceased animals should be 

necropsied to determine the cause of 

death. Disposal after necropsy must be 

done in accordance with local/federal 

laws. 

Otter SSP Quarantine Recommendations: All animals should undergo a 30-day quarantine stay at the 

receiving institution before incorporation into the rest of the collection. This allows time for the 

development of clinical signs of disease that may have been incubating before the animal was shipped. 

During the quarantine period, the animal should be observed for signs that may be associated with 

disease, such as sneezing, coughing, vomiting, diarrhea, ocular or nasal discharge, etc. Three fecal 

examinations for parasites should be performed and negative results obtained before release into their 

permanent enclosure. P. brasiliensis should be checked for Strongyloides species (Wünnemann 1990; 

C.Osmann, personal communication). The diet should be slowly adjusted over several weeks if there is to 

be a diet change. P. brasiliensis and A. cinereus should be housed in social groupings, as far as possible, 

during quarantine as extended periods alone can be detrimental to these social species causing 

development of stereotypical or self-mutilation behaviors such as pacing or over-grooming.



AZA Accreditation Standards and Related Policies 

See Appendix F for specific animal care and management recommendations for small carnivore 

quarantine, which are included in the AZA Accreditation Standards and Related Policies (2008). 

Quarantine Facilities: Ideally, quarantine facilities should be isolated from the risk of crosscontaminating 

other carnivores already in the collection. If this is not possible, different keepers should be 

used, or strict rules of personal hygiene should be adopted and resident animals should be cared for 

before quarantine animals. Balancing between the necessity of keeping the quarantine pen clean and the 

needs of the animals can be challenging. Many of the mustelid species do better when isolated in 

enclosures than when placed in hospital-type quarantine pens (Lewis 1995). If this is not practical or 

possible, a privacy box, climbing furniture, substrate suitable for rubbing/drying-off, and a pool or water 

tub suitable for swimming should be provided. Whatever type holding facility is used, care should be 

taken to ensure that otters cannot escape by climbing, digging, or chewing their way out. 

Quarantine Exams: Two quarantine exams are recommended for otters; one performed at the beginning 

of the quarantine period and one performed at the end. 

Initial exam: Veterinarians should visually inspect otters as soon as possible after they have arrived in 

quarantine. If a pre-shipment physical examination has not been done before the animal was transferred, 

it would be prudent to perform a complete examination during the first week of quarantine (see section 

6.2). 

Final exam: During the last week of quarantine, a thorough examination should be conducted as outlined 

in section 6.2. It is extremely important to take radiographs of the animal during this time even if they 

were done at the previous institution (see note on P. brasiliensis below). This gives the new institution its 

own baseline film from which to compare future radiographs. This is especially important since 

radiographic techniques vary from facility to facility. 

As giant otters are an endangered species that are very rare in zoos, the zoo animal population is 

highly valuable. The zoo veterinarian of the receiving institution should bear in mind that every anesthesia 

may be of high risk for each animal. Basic radiographs may be of importance from the medical point of 

view, but should only be taken if the otter is anesthetized for a special reason. Regular visual 

examinations of the otters’ health status during quarantine should be performed, as well as parasitological 

and microbiological testing of fecal samples (C.Osmann, personal communication). 

6.5 Preventive Medicine 

AZA Policies: AZA-accredited institutions should have an 

extensive veterinary program that must emphasize disease 

prevention (2.4.1). The American Association of Zoo 

Veterinarians (AAZV) has developed an outline of an effective 

preventative veterinary medicine program that should be 

implemented to ensure proactive veterinary care for all animals 

(www.aazv.org/associations/6442/files/zoo_aquarium_vet_med_g 

uidelines.pdf). 

As stated in Chapter 6.4, AZA institutions must have zoonotic 


(2.4.1) The veterinary care program must 

emphasize disease prevention. 


(11.1.2) Training and procedures must be 

in place regarding zoonotic diseases. 

disease prevention procedures and training protocols established to minimize the risk of transferable 

diseases (11.1.2) with all animals. Keepers should be designated to care for only healthy resident 

animals, however if they need to care for both quarantined and resident animals of the same class, they 

should care for the resident animals before caring for the quarantined animals. Care should be taken to 

ensure that these keepers are “decontaminated” before caring for the healthy resident animals again. 

Equipment used to feed, care for, and enrich the healthy resident animals should only be used with those 

animals. Standard institutional policy regarding disinfection of equipment used should be followed when 

working with otters. Care should be taken to ensure all disinfectant is washing from all surfaces before 

otters are reintroduced into an area. 

Also stated in Chapter 6.4, a tuberculin testing and surveillance program must be established for 

animal care staff, as appropriate, to protect the health of both staff and animals (11.1.3). Depending on 

the disease and history of the otters, testing protocols may vary from an initial quarantine test, to annual 

repetitions of diagnostic tests as determined by the veterinarian. To prevent specific disease

transmission, vaccinations should be updated as appropriate 

for otters. TB testing is not routinely done in otters. 

AZA Otter SSP Recommendations for Medical 

Management 




(11.1.3) A tuberculin testing and 

surveillance program must be established 

for appropriate staff in order to ensure the 

health of both the employees and the 

animal collection. 

Medical Examinations: It is recommended that all animals 

have at least a biennial examination and, if possible, an annual examination (see below for exception for 

P. brasiliensis), during which the following procedures are performed: 

• Transponders and/or tattoos should be checked and reapplied if they are not readable. 

• Baseline physiological parameters (e.g., heart rate, weight, body temperature, respiratory rate) 

should be obtained & recorded. 

• The oral cavity and all dentition should be examined. Teeth should be cleaned and polished if 

necessary. Any tooth that is fractured or in need of repair should be noted in the medical record 

and the condition corrected as soon as possible. 

• The reproductive tract should be evaluated. Care should be taken to record any changes in the 

external genitalia, such as vulvar swelling or discharge, testicular enlargement, and mammary 

gland changes. Contraceptive hormone implants also should be checked to make sure they are in 

place, and not causing any local irritation. 

• Radiographs taken to check for any abnormalities. If renal or cystic calculi are seen, then 

numbers, location, and approximate sizes should be noted in the records. 

• Blood collection done, and complete blood count and chemistry profile performed. For P. 

brasiliensis, blood samples can be taken from the V. cephalica antebrachii of the foreleg 

(C.Osmann, personal communication). 

• Blood serum frozen and banked when possible. 

• Animals housed outside in heartworm endemic areas should be checked for heartworm disease 

by performing a heartworm ELISA antigen test and the animal routinely given heartworm 

preventative treatment (see ‘parasite control’ section). 

• Urine collected whenever possible by cystocentesis for a complete urinalysis. For P. brasiliensis, 

rather than cystocentesis, which is associated with the risk of bacterial infection, it is possible to 

gain urine from an immobilized animal by manual squeezing of the caudal abdomen (female) or 

by catheterizing the urethra (C.Osmann, personal communication). 

• An annual fecal examination should be performed to check for internal parasites, and 

anthelmintics administered if necessary (see ‘parasite control’ section). For P. brasiliensis, 

biennial fecal examinations are recommended (C.Osmann, personal communication). 

• Vaccines updated if necessary (see ‘vaccination’ section). 

Anesthesia/immobilizations of giant otters should only be performed when there is a medical 

indication for the procedure. Preventive immobilizations are of high risk for the animals, and should be 

substituted with regular visual examinations and testing of fecal samples, vaccinations etc. Administration 

of transponders, examination of the oral cavity, blood sampling etc., should be completed only when 

immobilizations are necessary for medical reasons. Evaluation of the reproductive tract can be performed 

in animals that are regularly involved in a medical training program. In well-trained animals, sonography 

of the uterus may be possible, as well as the visual or palpable inspection of mammary gland and 

testicles. Contraceptive hormones should not be used in giant otters because of side effects such as 

endometritis and pyometra, and the potential result of future breeding inhibition (C.Osmann, personal 

communication)



Animal Records: Thorough and accurate medical records are essential to learn and understand more 

about the medical problems of species in ex-situ populations. Medical records should be systematic and 

entries should identify the history, physical findings, procedures performed, treatments administered, 

differential diagnosis, assessment, and future plans for treatment. A computerized medical record system, 

which can help track problems and be easily transmitted from one institution to the next, is extremely 

beneficial. The AZA Otter SSP encourages the use of Med ARKS (International Species Information 

System, 12101 Johnny Cake Ridge Road, Apple Valley, MN 55124, U.S.A.) as a universal medical record 

program. Many institutions already use this program, making it easy to transfer information between them. 

At this time the ZIMS product is being developed to replace current zoological record keeping systems 

and is considered a suitable substitute. 

Vaccination: The following vaccination schedule is recommended by the AZA Otter SSP Veterinary 

Advisor. Vaccination product recommendations are based on clinical experience (as of 2006) in most 

cases, and not necessarily on controlled scientific study. 

Canine distemper: Merial's new PureVax Ferret Distemper Vaccine currently on the market is a 

univalent, lyophilized product of a recombinant canary pox vector expressing canine distemper virus 

antigens. The vaccine cannot cause canine distemper under any circumstances, and its safety and 

immunogenicity have been demonstrated by vaccination and challenge tests. Otters should initially be 

given 1ml of reconstituted vaccine for a total of 2-3 injections at three-week intervals, followed by a yearly 

booster. This vaccine should be given IM instead of SQ in exotic carnivores for increased effectiveness. 

More information on PureVax Ferret Distemper Vaccine can be found at www.us.merial.com (Merial 

Ltd., 3239 Satellite Blvd., Duluth, GA 30096). An alternative vaccine that is available is Galaxy D 

(Schering-Plough Animal Heath Corporation, P.O. Box 3113, Omaha, NE 68103), a modified-live canine 

distemper vaccine of primate kidney tissue cell origin, Onderstepoort type. Safety and efficacy of canine 

distemper vaccinations in exotic species of carnivores have been problematic. Vaccine-induced 

distemper has occurred in a variety of mustelids using modified-live vaccine, and killed vaccines have not 

provided long-lived protection and are not commercially available. However, to date there have been no 

cases of vaccine induced distemper in otters given the Galaxy product, and excellent seroconversion 

following vaccination using this product has been documented in young N.A. river otters (K.Petrini, 

unpublished data, Petrini et al. 2001). The use of any modified-live canine distemper vaccine in exotic 

species should be done with care, especially with P. brasiliensis, young animals, and those that have not 

been vaccinated previously. The use of PureVax Ferret Distemper Vaccine is recommended where 

possible. 

Parvovirus: The efficacy of feline and canine parvovirus vaccines has not been proven in otters. Otters 

should initially be given 1ml of vaccine IM for a total of 2-3 injections at three-week intervals followed by a 

yearly booster. Parvocine (Biocor Animal Health Inc., 2720 North 84th Street, Omaha, NE 68134) is a 

killed univalent parvovirus vaccine that has been used in otters. Using a univalent product such as 

Parvocine reduces the risk of vaccine allergic reactions. 

Rabies: The efficacy of rabies vaccines has not been proven in Lontra canadensis or other exotic 

mustelids. Vaccinated otters that bite humans should not be considered protected from rabies. Only killed 

rabies products should be used in otters. One commonly used product is Imrab ® 3 (Merial Ltd., 3239 

Satellite Blvd., Duluth, GA 30096), which is a killed rabies vaccine that has been used extensively in small 

carnivores without apparent adverse effects. Otters should be given 1ml of vaccine IM once at 16 weeks 

of age followed by a yearly booster. PureVax Feline Rabies (Merial Ltd., 3239 Satellite Blvd., Duluth, GA 

30096) is a live canarypox vectored, non-adjuvanted recombinant rabies vaccine that is currently being 

used at some institutions for small carnivores. Dosage and route are the same as for Imrab ® 3, but this 

vaccine can be given once at age 8 weeks or older, then annually. 

Leptospirosis: The susceptibility of river otters to leptospirosis is debated in the literature, and the benefit 

of vaccination is unknown. Killed Leptospira bacterins are available and can be administered in areas 

where leptospirosis has been problematic. Initially two doses should be given at 3-4 week intervals. 

Vaccine efficacy and duration of immunity has not been studied in the otter and is an area where further 

study should be conducted. 

Parasite Control: Otters should have fecal examinations performed regularly (annual exams are 

advised). The frequency of these examinations depends on the incidence of parasitism in the geographic



region and the likelihood of exposure. Animals also should be screened for parasites before shipment and 

during quarantine. Fecal testing should include a direct smear examination and a fecal flotation, as well 

as sedimentation techniques, if possible. Baermann fecal examination techniques help identify certain 

parasites, such as lungworms, that are otherwise difficult to detect. 

Heartworm ELISA antigen tests should be conducted annually in animals exposed to mosquitoes in 

heartworm endemic areas and animals maintained on a heartworm preventative. External parasites (e.g., 

mites, fleas, ticks) can be detected during physical examinations. 

Dr. George Kollias, Cornell University School of Veterinary Medicine, states: “Dirofilaria immitis, the 

cause of heartworm disease in dogs, cats and some other carnivores, has been found in the hearts of 

otters in and from Louisiana. This filarial worm has to be differentiated from Dirofilaria lutrae, the 

microfilaria of which can be found in the blood and adults in the subcutaneous tissues and coelomic 

cavity of river otters. D. lutrae generally does not cause disease. Newly acquired otters should be 

screened for microfilaria (via the Knott's test on blood) and for adults, via the ELISA antigen test on 

serum. D. immitis can be differentiated from D. lutrae by the morphological appearance of the microfilaria 

and by the antigen test. Thoracic radiographs should also be taken as part of routine health screening 

and definitely if an otter is Knott's test positive and/or antigen positive”. See also Snyder et al. (1989), 

Neiffer et al. (2002), and Kiku et al. (2003) for reports of heartworm in otters. 

In heartworm endemic areas, otters can be given ivermectin (0.1mg/kg orally once/month year 

around) as a preventative. Although it is still uncertain whether or not D. immitis causes progressive 

heartworm disease, as in the dog and cat, prevention is safest approach. If used at the proper dose, 

ivermectin has proven safe in otters. Mortality has been associated with Melarsomine dihydrochloride 

administration to North American river otters and a red panda for heartworm disease (Neiffer et al. 2002). 

In another report of otter deaths after treatment with Melarsomine, adult heartworms were found in the 

hearts of three out of the four animals during necropsy (G. Kollias, personal communication). 

Parasite Testing: Recommendations and protocols for parasite testing in otters are provided in the 

following table: 

Table 5: Otter parasite testing protocols 

Parasite Testing protocol 

External parasites Regular inspections during any physical examinations 

Internal parasites Annual fecal examinations: direct smear, fecal flotation, & sedimentation or Baermann 

techniques. 

Pre-shipment fecal examinations: direct smear and flotation 

Quarantine fecal examination: 3 negative direct smear results & 3 negative fecal 

flotation results before release from quarantine. 

Heartworm ELISA antigen tests: conducted annually in animals exposed to 

mosquitoes in heartworm endemic areas (test will not detect all male infections or 

infections with < 3 female nematodes). If infection is suspected, positively identify the 

microfilaria as pathogenic before instituting treatment.



Parasite Treatment: The following table (Table 6) provides a list of anthelmintic products that have been 

used safely in a variety of mustelids: 

Table 6: Recommended anthelmintic treatments for otters 

Treatment Dose 

Fenbendazole 50mg/kg orally for 3-5 days. In P. brasiliensis there was a complete elimination of a 

Strongyloides spp. infestation after treatment with 10-20mg/kg over 3 days 

Pyrantel pamoate 10mg/kg orally 

Ivermectin 0.1mg/kg orally, once monthly for heartworm prevention 

Praziquantel 5mg/kg SC or orally 

0.2-0.4mg/kg subcutaneously or orally for treatment of intestinal nematodiasis 

(G.Kollias, personal communication) 

Medical Management of Neonates: Otter pups can develop health issues suddenly, and they should be 

carefully watched for any change in behavior. Some problems that have developed in young hand-reared 

pups are listed below with suggested first-step solutions or treatments, and neonatal examination and 

monitoring guidelines are also provided. See Table 7. 

Table 7: Neonatal examination & monitoring guidelines (from Read & Meier 1996) 

Vital signs Temperature, include activity level 

Pulse, rate and character 

Respiration, rate and character 

Organ systems --- 

Weight --- 

Hydration Skin tone and turgor 

Mucous membranes Color and capillary refill 

Vitality Response to stimulation, activity levels: type, frequency, duration 

Physical condition --- 

Laboratory values (optional) Complete blood count 

White blood cell count 

Serum chemistries, including blood glucose & blood urea nitrogen 

Urinalysis and urine specific gravity (recommended) 

Urination Frequency, amount, and character 

Defecation Frequency, amount, and character 

Condition of umbilicus --- 

Total fluid intake Amount in 24 hours 

Parenteral fluids, amount, frequency, and type 

Oral fluids, amount, frequency, type, nipple 

Housing temperature --- 

Dehydration/Emaciation: Give subcutaneous or oral (only if sucking well) electrolytes. Lactated Ringers 

Solution (LRS) with 2.5% dextrose or sodium chloride (0.8% NaCl) are recommended. Oral fluids are 

given at the dose of 5% body weight per feeding. The dose for subcutaneous fluids is determined by the 

level of dehydration and should be determined by a veterinarian.



P. brasiliensis: Parenteral volumen substitution can be made with Amynin ® (full electrolyte, glucose, 

amino acid solution offered by Merial), Ringer Lactate, and Glucose 5%, for a total dosage of about 10% 

of body weight/day. Oral fluids for children, such as Humana Elektrolyt ® , are a good option to use in otter 

pups (C.Osmann, personal communication). The AZA Otter SSP recommends veterinarians contact one 

of these companies for their advice on suitable products. Weak pups may be gavage-fed by someone 

experienced in inserting a tube into the stomach and injecting formula directly into the digestive tract. This 

is a risky endeavor, as the stomach-tube can be accidentally inserted into the trachea resulting in milk 

infusion directly into the lungs. In general, if pups are too weak to suckle, their gastrointestinal tracts are 

too compromised to digest food and they require immediate veterinary care. Administration of paramunity 

inducers, such as Zylexis ® (Pfizer - www.pfizer.com/pfizer/main.jsp), is recommended in weak and less 

vital cubs (C.Osmann, personal communication). 

Diarrhea/Constipation: Digestive upset is a common issue with hand-reared neonates, and may be 

associated with several factors (Meier 1985): inappropriate milk formula, feeding frequency, overfilling the 

stomach which can cause bloating, and rapid changes in the diet. When digestive upset occurs, 

characterized by diarrhea, bloating, inappetance, and/or extreme disorientation, it is recommended that 

one factor is analyzed and/or changed at a time. The veterinarian should be consulted immediately in the 

case of diarrhea, as the condition of very young animals can deteriorate rapidly. 

Diarrhea related to diet changes may be treated with Kaopectate ® with veterinary approval. It should 

be noted that Kaopectate ® now contains salicyclic acid (aspirin), as does Pepto-Bismol ® , and 

gastrointestinal bleeding may result from frequent doses. Persistent diarrhea, or loose stool accompanied 

with inappetance, requires continuous veterinary care. Bacterial infections or parasites, such as Coccidia 

may be the cause of the problem and require specific medication. Osmann (personal communication) 

recommends the administration of Lactobacillus spp. into the formula for P. brasiliensis pups with 

diarrhea or after antibiotic treatment. Veterinarians should consider this for all otter species. 

Constipation may be treated by diluting the formula to half-strength for 24 hours, and gradually 

increasing back to full-strength over a period of 48 hours. The pup also can be given oral electrolyte fluids 

at the rate of 5% body weight in between feedings and 1-2 times over a 24-hour period. The pup’s back 

end can be soaked for a few minutes in warm water (make sure to dry off completely) accompanied by 

gentle stimulation, but care should be exercised that the anal area is not irritated. 

Upper Respiratory Infections: Pups that have been eating normally and suddenly start chewing on the 

bottle or seem uninterested in the bottle may have an upper respiratory infection. They cannot nurse 

properly when congested. Upper respiratory infections need to be treated immediately. Newborn pups 

can die within 24 hours of the first symptom. Antibiotics should be started at the first sign of infection. 

Antibiotics can be given orally or injected. Care should be taken with the location of injections to avoid 

the sciatic nerve in their rear limbs (in two cases where limb mobility was affected due to injection site, the 

lameness/paralysis was resolved over time). Pups on antibiotics may also develop GI problems and/or 

get dehydrated, and this should be treated accordingly. Antibiotics that have been used successfully for 

upper respiratory infections are listed below. Antibiotics should not be given without consulting a 

veterinarian first. 

• Enrofloxacin: injectable at 5mg/kg BID IM 

• Amoxicillin: 20mg/kg BID PO 

• Amoxicillin (long-acting): 15mg/kg IM every 48 hours (P. brasiliensis) 

• Penicillin G Procaine: 40,000-44,000 IU/kg q24 hr IM 

• Chloramphenicol: administered orally at 30-50mg/kg/day (P. brasiliensis) 

• Trimethoprim/sulfonamide combination: given parenteral at 15mg/kg/day (P. brasiliensis – 

C.Osmann, personal communication) 

Bloat: Some otter pups have developed bloat. Care should be taken to ensure that there is no air in the 

formula or any leaks in the bottles. The amount of formula fed at each feeding should be re-evaluated as 

the pup may be receiving too much. Reducing the amount fed per feeding and adding another feeding 

should be considered. Watch for respiratory distress as respiration may become labored with severe 

abdominal distention. Treatment options for bloat include passing tubing to decompress, or the use of 

over-the-counter medication. Infant gas drops have been tried with no effect. Care should be taken with 

the use of certain gastric coating agents, such as bismuth subsalicylate (Pepto-Bismol ® ), as some 

ingredients may create more problems.



Fungal Infections: Caretakers should look for hair loss and discoloration of skin, and should pull hair 

samples and culture for fungus using commercially available fungal culture media. At first appearance, 

fungal infections can be treated with shampoos and creams, and shaving the affected areas can also 

help. Severe infections may need to be treated with oral/injectable medication. 

Parasites: Fecal samples should be taken regularly from otter pups (specifically hand-reared pups), even 

if they are negative. Pups should be dewormed as needed and treatment started immediately to avoid 

any weight loss. 

Bite/Puncture Wounds: Any bite or puncture wounds should first be cleaned and flushed with fluids, and 

then treated with topical antibiotic and systemic antibiotics if necessary. 

6.6 Capture, Restraint, and Immobilization 

The need for capturing, restraining and/or immobilizing an 

otter for normal or emergency husbandry procedures may be 

required. All capture equipment must be in good working order 

and available to authorized and trained animal care staff at all 

times (2.3.1). 


(2.3.1) Capture equipment must be in 

good working order and available to 

authorized, trained personnel at all times. 

It is recommended that anesthesia be given to otters intramuscularly (IM) in the cranial thigh 

(quadriceps), caudal thigh (semimembranosus-tendinosus), or paralumbar muscles (Spelman 1999). 

Animals should be kept as quiet as possible. Generally, restraint is accomplished using a net, squeeze 

cage, or capture box. The AZA Otter SSP recommends training animals to receive injections to minimize 

stress prior to all anesthesia events. Immobilization of P. brasiliensis using a blowpipe has proven to be 

relatively easy and minimizes stress to the animals involved. Osmann (personal communication) 

recommends darting the animal in the M. biceps femoris/semimembranosus/semitendinosus. A variety of 

agents have successfully been used in otter species for immobilization. These include Ketamine alone 

(not recommended), Ketamine with midazolam, Ketamine with diazepam, and Telazol ® . 

Otters have a large respiratory reserve, and so using gas induction chambers is often very time 

consuming (this can take up to 10 minutes in A. cinereus), but has been done successfully. Despite the 

method of induction, anesthesia can be maintained by intubating the animal and maintaining it on 

Isofluorane (Ohmeda Pharmaceutical Products Division Inc., P.O. Box 804, 110 Allen Rd., Liberty Corner, 

NJ 07938). Halothane (Fort Dodge, 9401 Indian Creek Parkway, Ste. 1500, Overland Park, KS 66210) is 

no longer recommended for use in otters as it may cause liver failure (G.Meyers, personal 

communication). Otters are relatively easy to intubate, and this method is preferred when it is necessary 

for an animal to be immobilized for a lengthy procedure (>30 minutes). 

Careful monitoring of anesthetic depth and vital signs is important in any immobilization. Body 

temperature, respiratory rate and depth, heart rate and rhythm, and mucous membrane color and refill 

time should be assessed frequently. Pulse oximetry sites include the tongue, the lip at the commissure of 

the mouth, or in the rectum. Oxygen supplementation should be available and administered when 

indicated. 

A. cinereus: For A. cinereus, ketamine hydrochloride can be used alone or in combination with midazolam 

hydrochloride (Versad ® , Roche Labs, 340 Kingsland St., Nutley, NJ 07110-1199) or diazepam to improve 

muscle relaxation (Petrini 1998). Telazol ® (Fort Dodge, 9401 Indian Creek Parkway, Ste. 1500, Overland 

Park, KS 66210) is another good immobilizing agent for this species. Generally, it provides smooth, rapid 

induction and recovery along with good muscle relaxation. Doses of Telazol ® required for adequate 

immobilization vary considerably between individuals. Ranges for some injectable drug combinations are 

listed below: 

• Telazol: 5.5-9.0mg/kg IM 

• Ketamine: 12-15mg/kg & midazolam: 0.5-0.75mg/kg IM 

• Ketamine: 9-12mg/kg & diazepam: 0.5-0.6mg/kg IM 

Muscle rigidity is common with these injectable drug combinations at the lower end of the dosages. 

Initial apnea and low oxygen saturation readings, as measured by pulse oximetry, often accompany 

higher doses. All three combinations produce a relatively short duration of anesthesia time, approximately 

15-30 minutes. Administering an additional 5mg/kg ketamine IM when needed can prolong anesthesia 

time. Alternatively, the animal can be intubated and maintained on gas anesthesia.



Combining ketamine with medetomidine hydrochloride (Domitor ® , Pfizer Animal Health, 812 

Springdale Dr., Exton, PA 19341) may provide a slightly longer duration of anesthesia and may give 

better myorelaxation; it also has the added advantage of being reversible with atipamezole hydrochloride 

(Antisedan, Pfizer Animal Health, 812 Springdale Dr., Exton, PA 19341). Vomiting may occur during 

induction, and initial apnea and low oxygen saturation readings are common. Supplemental oxygen 

should be available for administration if necessary. Dosages that have been used successfully are: 

• Ketamine: 4-5.5mg/kg & medetomidine: 0.04-0.055mg/kg IM; reversed with atipamezole: 0.200- 

0.275mg/kg IM 

L. canadensis: For short-term anesthesia (25-30 minutes) of L. canadensis, Spelman (1998) recommends 

the following: 

• Ketamine: 10mg/kg & midazolam: 0.25mg/kg 

• Ketamine: 2.5-3.5mg/kg, medetomidine: 0.025-0.035mg/kg & atipamezole: 0.125mg/kg 

(respiratory depression is more likely at higher dosages). 

• Telazol ® : 4mg/kg (Spelman 1998), 9mg/kg (Blundell et al. 1999; Bowyer et al. 2003), 8mg/kg 

(Petrini et al. 2001); reversed by Flumazenil: 0.08mg/kg to prevent a prolonged recovery time. 

• Ketamine: 10mg/kg. Muscle rigidity and variable duration should be expected. 

• Ketamine: 5-10mg/kg & diazepam: 0.5-1mg/kg. Prolonged recovery compared to ketamine with 

midazolam. 

P. brasiliensis: Due to their large size, a deep IM injection is recommended for good anesthesia. The 

breathing of the animals should be carefully monitored, and the temperature tested frequently to avoid 

hyperthermia (L.Spelman, personal communication, 2007). The following anesthesia protocols have been 

used with giant otters: 

• Ketamine at 7.5mg/kg (5-10mg) in combination with xylazine at 1.5mg (1-2mg/kg). Combining 

Ketamine with xylazine (Rompun ® 2%, BayerVital GmbH, 51368 Leverkusen) gives a short-term 

anesthesia with good muscle relaxation and analgesia. Xylazine may be reversed with 

atipamezole (Antisedan ® , Pfizer GmbH, Pfizerstraße 1, D-76139 Karlsruhe) (C.Osmann, personal 


• Give xylazine at 2.5mg/kg, wait 15 minutes and give ketamine at 2.5mg/kg; when done, reverse 

with yohimbine (L.Spelman, personal communication, 2007). 

• For a single injection, use medetomidine 0.030mg/kg and ketamine 3mg/kg, and reverse with 

atipamezole 0.125mg/kg. Although easier to use, this regimen can lead to poor breathing at the 

start of the procedure (L.Spelman, personal communication, 2007). 

• Supplemental oxygen should always be available for administration, if necessary. For longer 

procedures, animals should be maintained on Isofluorane. 

6.7 Management of Diseases, Disorders, Injuries and/or Isolation 

AZA Policies: AZA-accredited institutions should have an 

extensive veterinary program that manages animal diseases, 

disorders, or injuries and has the ability to isolate these animals 

in a hospital setting for treatment if necessary. Staff should be 

trained for meeting the animal’s dietary, husbandry, and 

enrichment needs, as well as in restraint techniques, and 

recognizing behavioral indicators animals may display when their 

health becomes compromised (2.4.2). Protocols should be 

established for reporting these observations to the veterinary 

department. Hospital facilities should have x-ray equipment or 

access to x-ray services (2.3.2), contain appropriate equipment 

and supplies on hand for treatment of diseases, disorders or 

injuries, and have staff available that are trained to address 

health issues, manage short and long term medical treatments 

and control for zoonotic disease transmission. 

AZA-accredited institutions must have a clear process for 


(2.4.2) Keepers should be trained to 

recognize abnormal behavior and clinical 

symptoms of illness and have knowledge 

of the diets, husbandry (including 

enrichment items and strategies), and 

restraint procedures required for the 

animals under their care. However, 

keepers should not evaluate illnesses nor 

prescribe treatment. 


(2.3.2) Hospital facilities should have xray 

equipment or have access to x-ray 

services. 

identifying and addressing animal welfare concerns within the institution (1.5.8) and should have an 

established Institutional Animal Welfare Committee (AWC). This process should identify the protocols 

needed for animal care staff members to communicate animal welfare questions or concerns to their

supervisors, their Institutional Animal Welfare Committee or if 

necessary, the AZA Animal Welfare Committee. Protocols 

should be in place to document the training of staff about otter 

welfare issues, identification of any otter welfare issues, 

coordination and implementation of appropriate responses to 

these issues, evaluation (and adjustment of these responses if 

necessary) of the outcome of these responses, and the 

dissemination of the knowledge gained from these issues. 

As stated in Chapter 6.4, all living animals will die at some 

point. As care givers for the animals residing in our zoos and 

aquariums, it is vital that we provide the best care possible for 

them until the time their health deteriorates to a point where 

euthanasia is the most humane treatment, or the animal dies 

on its own. Necropsies should be conducted on deceased 




(1.5.8) The institution must develop a 

clear process for identifying and 

addressing animal welfare concerns 

within the institution. 


(2.5.1) Deceased animals should be 

necropsied to determine the cause of 

death. Disposal after necropsy must be 

done in accordance with local/federal 

laws. 

otters to determine their cause of death and the subsequent disposal of the body must be done in 

accordance with any local, state, or federal laws (2.5.1). Necropsies should include a detailed external 

and internal gross morphological examination and representative tissue samples form the body organs 

should be submitted for histopathological examination. 

AZA Otter SSP Disease Management Recommendations: Little information on common diseases and 

disorders for A. capensis and L. maculicollis is available, and more research is required for these species. 

Urolithiasis is the most common illness, and renal calculi are the most frequent cause of death in A. 

cinereus. The cause and reversal of this condition is the subject of ongoing research. 

Poor Coat Quality: Otters are amphibious mammals reliant on trapping air within their coats rather than 

a layer of blubber for thermal insulation (Tarasoff 1974). Studies have shown that otter fur is far denser 

than that of other mammal species, with an average of 26,000 hairs/cm 2 (foot) to 165,000 hairs/cm 2 

(foreleg) (Weisel et al. 2005). Sea otter pelts are roughly twice as dense as the fur of a river otter, and the 

river otters’ fur is twice as dense as that of a mink (Weisel et al. 2005). 

Weisel et al. (2005) determined, via the use of scanning and polarizing light microscopy, that otter 

guard and underhairs are characterized by the presence of fins, petals, and grooves that allow adjacent 

hairs to fit together forming an interlocking structure. Trapped within this interlocking structure are bubbles 

of air forming an insulating layer between the skin and water. Air is trapped in the fur when the otter 

shakes upon emerging from the water, via piloerection of the hairs (including grooming and rubbing), and 

muscular pleating of the skin (Weisel et al. 2005). Thus, behavioral actions combined with the density and 

complexity of the underfur structure essentially prohibits water from touching the skin. Weisel et al. (2005) 

also determined that the outer and inner hairs of an otter’s coat are, “… hydrophobic due to the presence 

of a thin layer of body oil from the sebaceous glands of the otter.” 

This recent work documented that the long, outer hairs do guard the more fragile inner fur from 

damage and that they can become damaged, reducing their effectiveness. At this time it is not possible to 

say what damage is done to the otters’ guard hairs by gunite or other abrasive surfaces within ex-situ 

exhibits, but the AZA Otter SSP recommends that those surfaces be avoided in otter exhibits as far as 

possible. 

Poor coat quality and other factors can lead to pneumonia. Poor coat quality is of concern when its 

water repellency is affected. If water does not form droplets and cannot be easily shaken off the guard 

hairs (i.e., dark brown fur), the otters’ guard hairs clump together resulting in a coat that looks slick and 

saturated; this is an indication of poor coat quality. Poor quality leads to water penetrating the guard hairs 

and exposure of the under-fur (gray/white under coat), which can then become waterlogged. An otter in 

this condition may not swim in an effort to remain as dry as possible. If the otter does swim, and it cannot 

keep dry, its body temperature will drop rapidly leading to observable shivering, even during sleep. 

Enteritis can develop in cases of extreme chilling. If measures are not taken, death can follow in a matter 

of days through pneumonia and/or gastro-intestinal complications (Duplaix-Hall 1972). Insufficient land 

area compared to water area, and/or inappropriate enclosure substrates causing overly damp/wet 

conditions, were historically most often the reason for poor coat condition and the resulting health 

problems in river otters (Duplaix-Hall 1972, 1975). For the giant otter, this is still the most frequent cause 

of poor fur condition and related health problems; no other environmental or physical conditions have



been reported to cause these coat problems except in one case of an unrelated serious illness (Sykes- 

Gatz 2005, unpublished data). 

Common Disease Issues: Dr. Gwen Myers, AZA Otter SSP Veterinary Advisor conducted a review 

(G.Myers, personal communication) of all submitted necropsy reports for L. canadensis. Her findings 

(Table 8) indicate that the most frequent causes of L. canadensis deaths (excluding neonatal deaths).

Table 8: Common causes of deaths in L. Canadensis. 

Cause of death Causal factors 



Heart disease Heartworm/death from heartworm treatment; Acute myocarditis; 

Myofiber degeneration 

Renal failure Etiology unknown; Amyloidosis; Pyelonephritis 

Hepatic lipidosis 

Adenocarcinoma 

Transitional cell carcinoma (bladder) 

Peritonitis Secondary to intestinal perforation from foreign body; Secondary 

to GI perforation from ulcers 

Diarrhea Unknown etiology; Clostridial endotoxin; Helicobacter (also 

causing vomiting, weight loss); Salmonella 

Gastric dilatation with volvulus 

Pneumonia Often without identifying underlying cause 

Anesthetic death 

P. brasiliensis: Causes of death have included: leptospirosis, parvovirus, bronchopneumonia/pneumonia, 

internal bleeding, gastroenteritis, jejunum invagination, severe inbreeding resulting in inherited thyroid 

malfunction in pups, parental or older sibling neglect of, or mistreatment towards pups due to stress from 

human disturbances or inappropriate insufficient land vs. water area and/or enclosure substrate 

conditions, inappropriately conducted introductions of unfamiliar or temporarily separated otters, heart 

failure, kidney failure, pyrometra, and exposure to continually very damp or wet conditions (Osmann & 

Wisser 2000; Sykes-Gatz 2005, unpublished data; C.Osmann, personal communication). 

Illnesses seen in this species include skin lesions, particularly on the tail and hind legs. These often 

become infected with Staphylococcus spp. and typically respond well to topical and/or systemic 

antibiotics (C.Osmann, personal communication). Progressive walking difficulties involving the lower back 

or hind legs also are reported in this species, particularly in animals aged 4 years and over. Again, the 

causal agent appears to be continued overexposure to hard surfaces. Other physical problems caused by 

overexposure to hard or continuously wet or very damp conditions include foot pad abrasions, irritation of 

the foot’s webbing, and poor coat condition (Sykes-Gatz 2005). These conditions can be caused or made 

worse by exposure to coarse substrates. 

For additional information on how to deal with separated animals, see pages 29 (A. cinereus), 31 (P. 

brasiliensis) and 59. If one of the social otters (e.g. A. cinereus or P. brasiliensis) has to be separated for 

an extended period, it may be best to house them with another individual. Prolonged isolation for this 

species is not considered desirable. Extended separation of female L. canadensis may lead to 

reintroduction issues; the reintroduction of these animals should proceed cautiously and follow the 

standard introduction process (See Introduction/Reintroduction section). 

Useful Veterinary References: 

Baitchman, E.J. DVM, G.V. Kollias, DVM, PhD. 2000. Clinical anatomy of the North American river otter 

(Lontra canadensis). Journal of Zoo and Wildlife Medicine 31(4):473-483. 

Kimber, K. DVM, G.V. Kollias DVM, PhD. 2005. Evaluation of injury severity and hematologic and plasma 

biochemistry values for recently captured North American river otters (Lontra canadensis). 

Journal of Zoo and Wildlife Medicine 36(3):371-384. 

Neiffer, D.L. VMD, E.C. Klein VMD, P.P. Calle VMD, M. Linn DVM, S. P. Terrell DVM, R. L. Walker DVM, 

D. Todd AHT, C. C. Vice DVM, S. K. Marks DVM. 2002. Mortality associated with melarsomine



dihydrochloride administration in two North American river otters (Lontra canadensis) and a red 

panda (Ailurus fulgens fulgens). Journal of Zoo and Wildlife Medicine 33(3):242-248. 

Rothschild, D.M., T.L. Serfass, W.L. Seddon, L. Hegde, R.S. Fritz. 2008. Using fecal glucocorticoids to 

assess stress levels in captive river otters. Journal of Wildlife Management 72(1):138–142. 

Schettler, E. DVM, K. Müller DVM, G. Fritsch DVM, S. Kaiser DVM, L. Brunnberg, DR., K. Frölich PhD, 

DVM, G. Wibbelt DVM. 2007. Progressive ataxia in a captive North American river otter (Lontra 

canadensis) associated with brain stem spheroid formation. Journal of Zoo and Wildlife Medicine 

38(4):579-582. 

Swenson, J. DVM, J.W. Carpenter, DVM, K.S. Janardhan PhD, C. Ketz-Riley DVM, E. Brinkman DVM. 

Paresis in an Asian small-clawed otter (Aonyx cinereus) associated with vertebral and ischial 

osteolysis caused by a malignant lymphangiosarcoma. Journal of Zoo and Wildlife Medicine 

39(2):236-243. 

Tocidlowski, M.E. DVM, L.H. Spelman DVM, P.W. Sumner MS, M.K. Stoskopf DVM, PhD. 2000. 

Hematology and serum biochemistry parameters of North American river otters (Lontra 

canadensis). Journal of Zoo and Wildlife Medicine 31(4):484-490. 

Weber, M.A. DVM, M.Garner DVM. 2002. Cyanide toxicosis in Asian small-clawed otters (Amblonyx 

cinereus) secondary to ingestion of loquat (Eriobotrya japonica). 2002. Journal of Zoo and Wildlife 

Medicine 33(2):145-146. 

Weisel, J.W., C. Nagaswami, R.O. Peterson. 2005. River otter hair structure facilitates interlocking to 

impede penetration of water and allow trapping of air. Canadian Journal of Zoology 83(5):649- 

655.



Chapter 7. Reproduction 

7.1 Reproductive Physiology and Behavior 

It is important to have a comprehensive understanding of the reproductive physiology and behaviors 

of the otters in our care. This knowledge facilitates all aspects of reproduction, artificial insemination, 

birthing, rearing, and even contraception efforts that AZA-accredited zoos and aquariums strive to 

achieve. 

A. cinereus: These otters are non-seasonal and thought possibly to be spontaneous ovulators (Bateman 

et al. 2009). The estrous cycle lasts 30-37 days, with breeding occurring year round. Some facilities 

report this cycle extending to every few months with older animals. Estrus lasts from 1-13 days. 

Behavioral signs of the onset of estrus may include increased rubbing and marking. Sexual behavior has 

been observed in pups as young as 6 months, with breeding behavior having been noted in animals 

(males and females) as young as 1½ years. Successful breeding has been reported for 2.1-year-old 

females and 2.8-year-old males. There do not appear to be any significant environmental cues that are 

involved with the onset of estrus. Breeding pairs have been introduced at various ages and have been 

together for varying lengths of time before successful breeding occurs. It has been reported that pups 

from previous litters have interfered with copulation, but their presence had no bearing in any other way 

(Lombardi et al. 1998). 

Recent work has shed light on litter intervals; Bateman et al. (2009) report: “In one female having 

three consecutive pregnancies during [their] study, the interval between the first parturition and 

subsequent progesterone increases owing to the next pregnancy was 169.25±11.15 days. This female 

was observed nursing her pups from the first pregnancy for the first 122 days of this intergestational time 

period.” 

Breeding pairs need to establish a bond for successful reproduction. The male pursues the female in 

courtship and most breeding occurs in shallow water. A single copulation can last from 5-25 minutes. 

Courtship behavior has been recorded from 1-3 days, at one-month intervals. Gestation is roughly 60-74 

days (67-77 range, average 71.17±1.49 days reported by Bateman et al. 2009). 

Pseudopregnancies do occur in this species, including in females housed in single sex groups (5 of 6 

females housed together exhibited pseudopregnancy) (Bateman et al. 2009). Bateman et al. (2009) 

report: “…a mean duration of pseudopregnancy of 72.45±1.37 days (range: 62–84 days). The average 

interval length between sequential pseudopregnancies and/or pregnancies was 39.86±3.86 days (range: 

17–92 days) in paired females and 134.50±48.94 days (range: 62–279 days) in the single gender group.” 

The sire plays a very active role in rearing the pups and should not be removed prior to their birth. 

Male behaviors include nest building, carrying pups, and bringing food to the pups during weaning.



Table 9: Ex-situ population breeding parameters of Aonyx cinereus in North American zoological facilities 1980's and 

1990's (Reed-Smith & Polechla 2002, Bateman et al. 2009) 

Aonyx cinereus 

Estrus cycle 30-37 days. Polyestrual with breeding occurring year around 

Estrus length 

1-10 days 

Ovulation 

Likely spontaneous ovulators (Bateman et al. 2009) 

Copulation frequency Several times a day. 

Copulation duration 1-30 minutes, varied. 

Copulation position Dorsal/Ventral most common, also ventro/ventral. 

Copulation location In the water and on land 

Copulation initiated by Varies amongst groups; in some it is initiated by male only, in others both initiate. 

Age at 1 st breeding Unknown 

Breeding behavior Increased rubbing, marking. 

Gestation Gestation ranges between 67 and 74 days; pseudopregnancies lasted 62 – 84 

days (Bateman et al. 2009). No delayed implantation (Bateman et al. 2009) 

Pair management Most facilities started out with a pair when 1st litter born. Pair left together all of the 

time 

Group management Some reports of harassment of new pups by older pups (too much play), one report 

reported cannibalism of pups by dam. Most leave all animals together. 

Signs of parturition Some weight gain, more time spent in nestbox. 

Pupping boxes Wooden boxes, hollows under logs & burlap bags have been used. 

Contraception MGA implants in females. 

A. capensis: This species does not appear to have a specific reproductive season (Mead 1989). 

Breeding in the northern hemisphere has been observed in November, January, March, and April 

(R.Meyerson, personal communication), with pups born in January-March and June-September 

(R.Meyerson, personal communication). Gestation length ranges from 63-80 days depending upon the 

source (Estes 1989; Reed-Smith & Polechla 2002; R.Meyerson, personal communication). In one ex-situ 

population breeding situation, receptivity by the female lasted one day (Personal communication); in 

another, breeding occurred for 2-3 days (R.Meyerson, personal communication). Generally, 1-2 days 

before a female is receptive, the male will start following her around. All ex-situ population pairs have 

shown an increase in the level of interactive play behavior for several days before and after breeding. All 

observed copulations have occurred in the water.



Table 10: Ex-situ population breeding parameters of Aonyx capensis in North American zoological facilities 1980's 

and 1990's (Reed-Smith & Polechla 2002) 

Aonyx capensis 

Estrus cycle Breeding occurred in Jan, Apr, Nov & Dec for 4 litters produced. 

Estrus length Peak receptivity lasted 1 day; day before & after consisted of play & close 

following 

Copulation frequency Several times during the 2 nd day of breeding behavior 

Copulation duration Not documented. 

Copulation position Not documented. 

Copulation location In the water. 

Copulation initiated by Male, females would only cooperate for 1 day. 

Age at 1 st breeding Males from 2 yrs.10 mos.; Females from 4 yrs. 2 mos. 

Breeding behavior Females fought when one was in heat. Signs of estrus shown by male behavior 

Gestation 80, 103 days 

Pair management Breeding was opportunistic. Male kept away from pups by female or separated 1 

week prior to expected parturition. 

Group management 1.2 housed together on exhibit during the day; 0.2 given 2 dens at night 

Signs of parturition Females gained weight particularly in fold between foreleg and body. 

Pupping boxes Females did not want bedding in their boxes. Given only one box, no problems. 

Contraception 

Two males have been vasectomized due to small gene pool. Females medicated 

for contraception. 

L. canadensis: These otters are seasonal breeders. Females mature reproductively as early as 12-15 

months (rare reports of successful breeding at this age) to two years of age (typical). They are believed to 

be induced ovulators and experience delayed implantation (Chanin 1985; Reed-Smith 2001, personal 

observation). Recent evidence suggests that this species also may be capable of spontaneous ovulation 

(Bateman et al. 2009). 

There is evidence that breeding season varies somewhat with latitude (Reed-Smith 1994, 2001; 

Bateman et al. 2005, 2009) and also may be influenced by seasonal availability of food resources (Crait 

et al. 2006); however, the authors of the one study (Crait et al. 2006) speculating on the influence of food 

availability acknowledge there could have been other things occurring, and their sample size was small. 

In general, breeding occurs in late spring (March-June) at northern latitudes and between November- 

February at more southern latitudes, with a gradient in between (Reed-Smith 2001). The estrus period 

lasts approximately 42-46 days, unless mating occurs (Chanin 1985). Bateman et al. (2009) found that 

“…peaks in fecal estrogen values occurred only during the defined breeding season from December to 

March”. They also report, “…the estrus phase of their cycles [N=11] was observed just once per year with 

an average duration of 15.33±1.98 days (range: 6–54 days). The average duration of estrus elevation 

coincident with observed breeding (n=4) was 22.00±1.22 days (range: 19–24 days).” During this time, 

observations of ex-situ populations suggest peaks of maximum receptivity are roughly 3-6 days apart with 

intervals of only mild receptivity during which the female may completely reject the male (Liers 1951; 

Reed-Smith 2001). The work done by Bateman et al. calls into question the estrus duration of 42 to 46 

days traditionally cited; this is an area that should be researched further. Worth noting is the slightly 

longer estrus (21-23 days) reported in breeding versus non-breeding females (14-17 days); also the 

widely varying range of estrus duration (6-54 days) reported in the Bateman et al. study. 

More recently, Bateman et al. (2009) have reported some additional interesting results from fecal 

hormone studies: 

“In the observed pregnancies and pseudopregnancies (n= 12), the date of initial 

progesterone increase ranged from September 4 to January 14, and the timing was 

not correlated (r=0.53, P>0.05) with the female’s geographic latitude (range: 27– 

41°N) at the time of the pregnancy or pseudopregnancy. However, the date of the



progesterone increase was correlated (r=0.66, P



Consequently, a possibility exists that some animals moved between widely varying latitudes may be 

physiologically out of synchrony and would require at least one breeding season to adapt physiologically 

to their new environmental cues, which are important for signaling the start of breeding season. This 

should be factored in when making transfer recommendations, but should not limit transfer options when 

creating new breeding pairs. The North American River Otter Husbandry Notebook (Reed-Smith 2001) 

provides greater detail on the breeding strategy and reproductive physiology of this species, and the AZA 

Otter SSP reproductive advisor (Helen Bateman, C.R.E.W., Cincinnati Zoo) is involved in on-going 

research. 

In both NARO and ASCO, additional research is needed to improve endocrine monitoring of estrogen 

metabolites to further address these questions about ovarian cyclicity and ovulatory mechanisms.



Table 11: Ex-situ population breeding parameters of Lutra canadensis in North American zoological facilities 1980's 

and 1990's (Reed-Smith & Polechla 2002, Bateman et al. 2009) 

Lutra canadensis 

Estrus cycle Monoestrus; can occur Nov-Jun based on latitude 

Post-partum elevations in estradiol levels occur 2 – 38 (ave. 19 ± 8.06) days after 

parturition (Bateman et al. 2009) 

Estrus length 

42-46 days unless mating occurs; “…the estrus phase of their cycles [N=11] was 

observed just once per year with an average duration of 15.33±1.98 days (range: 

6–54 days). The average duration of estrus elevation coincident with observed 

breeding (n=4) was 22.00±1.22 days (range: 19–24 days).” (Bateman et al. 2009) 

Receptivity peaks roughly 6 days apart have been reported but not reflected in 

Bateman study. 

Unclear if they are induced ovulators (Bateman et al. 2009), but suspected; may 

Ovulation 

also ovulate spontaneously (Bateman et al. 2009) 

Copulation frequency Several times a day 

Copulation duration 20-45 minutes, varied. One >60 min. reported 

Copulation position Dorsal/ventral most common, also ventro/ventral. 

Copulation location Most frequently in the water, also seen on land 

Copulation initiated by Both. Female advertises, cooperates only when she is ready. She may initiate 

with invitations to play chase. 

Age at 1 st breeding Sexually mature by 2 yrs. Several 2 yr. old males & a 1.5 yr. old female have bred 

successfully. 

Breeding behavior Female may rub, mark or vocalize to advertise; male/female may initiate w/ play, 

chase, splashing, genital sniffing, or “butterfly stroke”. 

Gestation 

Total gestation: 332-370 days, documented for 12 litters. 285-380 (Liers 1951); 

302-351 (ave. 333.3 ±15.7) days (Bateman et al. 2009) 

Actual gestation: 68 – 73 (ave. 71.67±1.48) days (Bateman et al. 2009) 

Pseudopregnancies seen with and without breeding and does not always result 

Pseudopregnancy when breedings are unsuccessful. The period of elevated progesterone ranges 

from 68 to 72 days as in true pregnancies. (Bateman et al. 2009) 

Pair management Most facilities separate the male from the female, for his safety. A few leave the 

male in exhibit with female and she keeps him away from pups until they can 

swim well. One facility offered pair selection; females showed preference for 

certain males. 

Group management Sire can be reintroduced to female and pups after pups are swimming well. 

Generally done at about 3-6 months. 

Signs of parturition Females may show visible weight gain, teats may show through coat, increased 

nesting behavior, change in attitude to keeper &/or male. She may go off food as 

parturition nears. 

Pupping boxes Pupping boxes should be filled with dry bedding (straw or hay). A choice of 

birthing boxes should be available. 

Contraception 

MGA implants and PZP treatment in females. A few males have been neutered. 

L. maculicollis: Schollhamer (1987) reported that females came into estrus for the first time at about two 

years of age, but females typically do not conceive until they are three years old. Cycles vary between 

individuals, but average about 45 days, and estrus generally lasts 5-7 days. There is a postpartum estrus 

2-3 weeks after parturition if pups are pulled or die soon after birth. 

Males may attempt breeding at 1-2 years of age, but typically are not successful until they are 2-3 

years old (Schollhamer 1987). Mating occurs in the water (Schollhamer 1987; R.Willison, personal 

communication) and involves the male neck biting the female and clasping her with his fore and hind 

limbs. Copulatory bouts may last up to 45 minutes, generally occur repeatedly over the course of several 

days (R.Willison, personal communication), and at any time during the day or night. There are no



vocalizations associated with copulation in this species in zoos (R.Willison, personal communication) or in 

the wild (Reed-Smith in prep.). The rate of conception is increased if the male breeds the female for the 

entire 5-7 day cycle of a typical female; the conception rate is poor if the male only breeds the female for 

2-3 days (Schollhamer 1987). 

Gestation is roughly 60-63 days (R.Willison, personal communication). Ex-situ population births in 

North America have occurred in January/February and April (R.Willison, personal communication). Births 

in the wild were recorded during September, based on one year of observation in Tanzania (Proctor 

1963); ongoing research in Tanzania substantiates this observation with breeding observed twice in June 

(2006 and 2007), which would result in late August to mid-September births. However, work by Bateman 

et al. (2009) and observations of half-grown animals throughout the year in Tanzania indicate that births 

may occur anytime with a peak during August/September. Further research is required. 

P. brasiliensis: Estrus generally occurs every three months, typically lasting 5-7 days with a range of 1- 

11 days (Autuori & Deutsch 1977; Trebbau 1978; Hagenbeck & Wünnemann 1992; Wünnemann 1995b; 

Marcato de Oliveira 1995; Corredor & Muñoz 2004; Sykes-Gatz 2005, 1999-2006). The pair will begin 

exhibiting an increase in rough play and chasing behaviors a few days prior to breeding. These behaviors 

continue throughout the estrus period (S.Sykes-Gatz, personal observation). Copulation generally takes 

place in the water, but also may occur on land. The copulatory act is typically repeated several times a 

day over the course of 5-7 days and may last from 30 seconds to 30 minutes or more (Hagenbeck & 

Wünnemann 1992; V.Gatz, personal observation; S.Sykes-Gatz, personal observation). 

There can be an estrus 5-7 days postpartum/post-loss of a litter that lasts for 3-5 days (Hagenbeck & 

Wünnemann 1992; Wünnemann 1995b). Delayed implantation occurs in zoos (Flügger 1997; Corredor & 

Muñoz 2004; Sykes-Gatz 2005, 1999-2006; V.Gatz, personal observation). Generally, this species 

produces one litter annually in the wild. In zoos, false or pseudo pregnancies are not uncommon in this 

species (Sykes-Gatz 2005). Gestation ranges from 64-71 days, and in one case a gestation of 77 days 

occurred in a 9-year-old, although this was unusually long for this female (Autuori & Deutsch 1977; 

Trebbau 1978; Hagenbeck & Wünnemann 1992; Wünnemann 1995a,b; Corredor & Muñoz 2004; Sykes- 

Gatz 2005; V.Gatz, personal communication). Often the female’s mammary glands become enlarged 30 

days prior to parturition and the vulva may become swollen about 14 days prior. In zoos and aquariums, 

inter-birth intervals of 63 days, 74 days, and 94-103 days have been recorded in cases of pairs that lost 

litters at birth (Hagenbeck & Wünnemann 1992; Wünnemann 1995b). 

Sexual maturity is reached at roughly 2 years of age. Ex-situ population records show that at 2 years 

and 5 months, females can come into their first estrus, mate, and bear a litter at the age of 2 years 7 

months. Males can mate at 2.5 years of age, with their first litter born when they are 2 years and 8 

months. Due to limited records, it is not known if this is the earliest age at which giant otters can become 

sexually mature and bear litters. There is some indication that at least some giant otter females, from the 

age of 10-11 years, may experience a slowing or end to their reproductive capabilities. Females of this 

age may alternatively experience health problems or other difficulties during gestation and parturition. 

Whether this is due to their advancing age or a high number of previous litters is unknown. An 18 years 

and 9 month old male is the oldest successful sire on record (Sykes-Gatz 2005 & 1999-2006, V.Gatz, 

personal communication). 

The International Giant Otter Studbook Husbandry and Management Information and Guidelines 

(Sykes-Gatz 2005) should be consulted for greater detail on management of this species, particularly 

their requirement for isolation and pupping den specifications. 

Separation of Sexes/Conspecifics: If it is necessary to separate animals for reasons associated with 

reproduction (e.g., to promote or prevent it), a holding area connected to the exhibit is recommended; 

ideally this should include a pool with clean water available at all times, proper lighting, a sleeping or den 

box, enough floor space for grooming and drying areas, and at least one nest box that is heavily bedded 

to allow excess moisture to be removed from the animals’ coats (Lombardi et al. 1998; Reed-Smith 2001). 

Holding pens should have non-climbable sides. If chain-link barriers are present, the sides should be 

covered with lexan or similar material to prevent animals from climbing too high and falling. See speciesspecific 

recommendations below. 

A. cinereus: Females become very aggressive prior to parturition and remain so post-parturition. It is not 

necessary to separate them from the male or older siblings. It is necessary to provide multiple nest boxes. 

The sire plays a very active role in rearing the pups and should not be removed prior to their birth. Male



behaviors include nest building, carrying pups, and bringing food to the pups during weaning. Access to 

pools and water sources should be strictly monitored to prevent newborns from drowning. 

A. capensis: Pregnant females should be offered nest box choices and separated from the male to give 

birth (R.Meyerson, unpublished data). To date, records indicate two institutions have successfully bred 

this species in North America. One facility had one male with two females; both females bred and 

produced offspring. The other facility housed a pair. The animals were housed together 24 hours a day; 

females were separated to give birth at both facilities (Reed-Smith & Polechla 2002; R.Meyerson, 

personal communication). The male can be reintroduced to the female and pups when they are swimming 

well. 

L. canadensis: There seems to be a mate preference for breeding, with some females showing a definite 

preference for particular male and lack of interest in others when they have a choice. Some successful 

zoos separate pairs for several months prior to the breeding season, introducing them every few days 

once the female’s estrus begins. Others offer multiple mate selections to the females, and others have 

been successful keeping single pairs together year around. An extensive ex-situ population study 

(N=13.14 animals) attempted to determine breeding associated behaviors in zoos and aquariums across 

the species over two years. The study’s behavioral results show an increase in pair association, mutual 

grooming, and extended copulation in those pairs that reproduced successfully (N=3), but was 

inconclusive otherwise (J.Reed-Smith, data in preparation). 

In the wild, males do not participate in pup rearing (Melquist & Hornocker 1983; Rock et al. 1994). In 

zoos and aquariums, parturient females should be given privacy (particularly for primiparous females) and 

nest box choices supplied with plenty of dry bedding (all females). Males have been successfully left in 

the exhibit with parturient females in large exhibits that provide numerous visual barriers and allow the 

male to stay out of the female’s sight. In all other cases, the male and female should be separated prior to 

the birth to prevent injury to the male or neglect of the pups by the female. In multi-female groups, other 

females may also need to be separated from the parturient female. When separated, the male or non 

parturient female should not be required to pass the parturient female’s den to enter the exhibit; if this 

cannot be done, the other animals should be removed entirely from the exhibit or the female sequestered 

away until she deems it time for the pups to meet the male (see below). Males can be reintroduced to the 

female and pups once they are swimming proficiently, as early as 60-75 days or more typically by 80 to 

90 days (Reed-Smith 2001). 

Actual gestation is calculated at roughly 68-73 days (Bateman et al. 2009.); pair separation should 

occur either when the female becomes aggressive towards the male or roughly 10-14 days prior to 

anticipated parturition date. Due to this species’ delayed implantation and total gestation time of >10 

months, it is often difficult to anticipate delivery date, particularly for primiparous females. In these cases, 

staff should base their management decisions on the female’s behavior. If she becomes aggressive to the 

male or other exhibit mates, begins to show excessive nesting behavior, or spends increasing amounts of 

time in her nest box, the pair should be separated. It is important to remember that the female should be 

monitored for health issues during this time, as these behaviors also can be signs of illness. 

Management change should be scheduled so that they do not interfere with the birth and rearing of 

the pups. Any modifications to the exhibit should be finished several months prior to possible pupping 

season. Denning/nest box choices should be introduced at least one month prior to possible pupping 

season. Changes in management routines, e.g., closing the female in holding at night, closing her in 

holding alone, etc. should be introduced to the female at least a month prior to the possible parturition 

period to allow her time to become comfortable with the new routine. If the male will have to be removed 

from the exhibit entirely, this should be done several weeks prior to possible parturition to allow the 

female a period of adjustment. 

Generally, the best way to handle pair separation is setting up the female in off-exhibit holding 

(providing there is adequate space). Once the pups are old enough to begin swimming lessons (some 

females begin this instruction as early as 30 days, more typically at 40-50 days), the family group and the 

male can be alternated on exhibit. When the pups are swimming well, after about three months, the male 

can be introduced to the family group. This should be handled, as with any introduction, via olfactory, 

visual, and then physical introduction to the female alone first, and under controlled circumstances as far 

as possible (see Introduction/Reintroduction).



L. maculicollis: Females should be separated from the male at about gestation day 55 (gestation ranges 

from 60-63+ days calculating from the day of last observed breeding) (Schollhamer 1987), or when she 

shows signs of aggression towards the male (R.Willison, personal communication). Males should be 

separated from a pregnant/nursing female unless the exhibit is large enough for him to stay out of her 

line-of-sight. When separated, he should not be required to pass the female’s den to enter the exhibit; if 

this cannot be done, he should be removed entirely from the exhibit or the female sequestered away until 

she deems it time for the pups to meet the male. Females with pups are more of a danger to the male 

than typical males are to the pups. Males can be reintroduced to the female and pups once they are 

swimming proficiently and eating on their own, typically when the pups are roughly four months of age 

(R.Willison, personal communication). 

Females should always be given a choice of denning sites with bedding provided for them to use if it 

is wanted. Schollhamer (1987) states that at Institutin A female spotted-necked otters did not use bedding 

of any kind. Kruuk (1995) references the presence of soft, leafy substrate or pebbles in most dens he or 

other researchers located. Brookfield Zoo used a nest box made from molded plastic and fiberglass 

measuring 68.6cm long x 51cm wide x 38.1cm high (27" x 20" x 15") with holes drilled in the bottom for 

drainage. The box was placed 4cm (1.5in) off the floor, and was accessed via a drop-guillotine door 

25.4cm high x 20.3cm wide (10" x 8") (Schollhamer 1987). 

P. brasiliensis: This species lives in family groups with pairs and older offspring jointly raising new pups. 

Therefore, a pair should never be separated during pregnancy or pup rearing. Typically, animals should 

not be separated from the family group unless health problems, change in social status, or family friction 

develops. Removal of any member of a group during pup-rearing, or close to parturition, will likely cause 

litter loss. Animals separated for extended periods should be put through a standard introduction (see 

Chapter 4, section 4.3), including visual, acoustic, and olfactory contact at first, and then physical contact 

(Sykes-Gatz 2005). Even a few days of separation have been known to be long enough to cause 

difficulty, such as serious fighting, when reintroduction was attempted (K.Lengel, personal 


Secondary accommodations should be provided for giant otters to allow for the temporary separation 

of family members if needed. These secondary enclosures should provide husbandry conditions similar to 

primary enclosures. 

Nursery Groups: Typically, nursery groups of neonates are not seen in river otter species. The following 

species-specific information is available: 

A. cinereus: All otters of a family group take an active role in caring for the young. It is not uncommon for 

the sire and older offspring to be involved in all behavioral activities of the mother and her newborns. 

A. capensis: Nursery groups are not typical for this species. 

L. canadensis: Nursery groups are not typical for this species. Helper otters have been reported from the 

wild. In these cases, a female with partially grown pups is accompanied by another adult female (Rock et 

al. 1994; R.Landis, personal communication). There are reports of two adults with young animals, but the 

relational composition of these groups is unknown (Reed-Smith 2001). 

L. maculicollis: Nursery groups are not reported for this species, but further research is needed. 

Adolescent groups (roughly one year or older) have been reported (Reed-Smith in prep.). 

P. brasiliensis: As with the Asian small-clawed otter, generational groups are typical in the wild; true 

nursery groups are not reported for either species. 

Separation of Mother and Offspring: The timing of mother-offspring separations can have long-term 

effects on the development of otter pups and on the reproductive success of adults. The following 

species-specific information is available: 

A. cinereus: Adolescents are not forced from the group. In zoos and aquariums, it is necessary to remove 

older offspring, as the group size can become quite large in a year, leading to aggression resulting from 

over-crowding. Typically, the age at which older pups should be removed varies with the size of the 

exhibit and compatibility of the group. 

A. capensis: The timing of emigration is unknown in the wild, but emigration of sub-adults at some point is 

presumed.



L. canadensis: Pups can be removed from the dam when weaned, if absolutely necessary. It is preferable 

that they be left with the family group until they are at least 8-9 months of age (Reed-Smith 2001) or six 

months old at a minimum. In the wild, pups will generally leave the female when they are 9 months to 

over one year of age (Melquist & Hornocker 1983; Melquist & Dronkert 1987). 

L. maculicollis: No specific information on emigration is available, but pups should be left in the family 

group for at least 6-9 months and be removed before reaching sexual maturity. Roving groups of what 

appear to be young animals, possibly dispersing, and pups remaining with their mothers (or at least 

utilizing the same core area simultaneously for one year) has been reported in the wild (Reed-Smith in 

prep). 

P. brasiliensis: Pups from previous litters up to the age of about two years generally stay with the family 

group (Schenck & Staib 1994); after this time, they emigrate to set up a new family group. In the wild, 

Duplaix (2002) reports that sub-adults may leave the family group after 2 years, before birth of the next 

litter, or be pushed out by the adults with a fight. Staib (2002) found that animals dispersed at the age of 

2-3 years, and separations were gradual, without aggressive behavior. In zoos and aquariums, offspring 

should be left with the parents for at least the first 6-12 months of life, but preferably they should be left 

together until they reach sexual maturity at approximately 2 years of age (Sykes-Gatz 2005, 1999-2006; 

Corredor & Muñoz 2004; G.Corredor, personal communication; V.Gatz, personal communication). This 

provides a more natural social structure, and allows older siblings to gain experience helping to rear pups, 

which is highly beneficial towards developing their future parenting skills. Caution should be taken, as in 

one case three ex-situ population-born otters between the ages of 6.5-8.5 months were suspected of 

competing for milk with their younger siblings. This behavior persisted over a two-month period, causing 

the death of a younger sibling and the necessary removal of the emaciated survivors for hand- rearing 

(Flügger 1997). In two other cases, sub-adults were suspected to have caused litter loss because of their 

over-zealous play with, and attention to, their younger siblings (Flügger 1997; G.Corredor, personal 

communication). However, experience has shown that removal of any member of a giant otter group 

during pup-rearing or close to parturition will likely cause litter loss due to the excessive stress caused to 

the parents by this human disturbance and unnatural social structure change (Flügger 1997; Sykes-Gatz 

2005, 1999-2006). If it is necessary to remove a group member from a breeding pair, it should be done 

when the mother is not pregnant or at the latest in the early stages of pregnancy. Pups younger than 6 

months of age should not be removed. 

Reproductive Hormone Tracking: Research utilizing techniques to identify reproductive state in these 

species is ongoing. At this time, it appears that ELISA protocols for testing hormonal secretions in fecal 

samples is successful in determining pregnancy in Asian small-clawed and North American river otters 

(Bateman et al. 2005, 2009). The reproductive physiology advisor for the AZA Otter SSP should be 

contacted for more information. 

Pseudopregnancy has been reported for most otter species and is an area that requires further 

research. For information on the status of current research into this and other reproductive physiology, 

behavioral, and health issues, contact the current AZA Otter SSP Chair for the most recent information. 

Facilities for Reproduction: All expectant females should be provided with nest box choices that are 

located away from pools, and these should be well stocked with dry bedding. The size of these dens 

should allow ample room for bedding, pups, and for the female to turn around (A. capensis, L. 

canadensis, and L. maculicollis). Highly social species (A. cinereus and P. brasiliensis) should be 

provided with a nest box or pupping den that allows enough room for the entire group. See speciesspecific 

information below and Sykes-Gatz for dimensions and recommendations on pupping dens and 

nest boxes. 

A. capensis: Nest boxes 8-10cm (3-4in) wider and taller than those used for L. canadensis (see below) 

are suitable for this species. Nest box choices and plenty of bedding should be provided 2-3 weeks 

before expected parturition date to allow the female to become comfortable with them. At this stage, 

females show a weight gain in the axillary region (R.Meyerson, personal communication). Some females 

prefer to pup without the bedding, and will remove it from their nest box; in these cases, allow the female 

her choice.



L. canadensis: Due to delayed implantation (also known as embryonic diapause), it is difficult to 

determine when a female is near parturition; therefore, close attention should be paid to her behavior 

changes, appetite, and physical appearance. These may include, but are not limited to: aggression 

towards exhibit mates or keeper staff, refusal to leave holding or her den, increased or decreased 

appetite, obvious teat development, slow movements, more frequent floating in the pool, and lethargy. 

Parturition boxes should be at least 50.8cm long x 45.72cm wide x 38.1cm high (20" x 18" x 15"), be 

large enough for an adult animal to move around in, and large enough to accommodate the pups. Slightly 

smaller boxes with entrance foyers have been used successfully. This box type allows the female to be 

secluded in a location near the pups but not actually with them. Females should be offered denning 

choices for parturition and den choices of different sizes to allow for the growth of the pups. 

Dimensions for a sample nest box are as follows: total width = 68.58cm (27in), chamber width = 

48.26cm (19in), chamber depth = 45.72cm (18in), entrance foyer = 20.32cm (8in), entrance diameter = 

16.5cm (6.5in), and 26.67-38cm (10.5-16in) high, with a 16.5cm (6.5in) height at the end of the ramp, and 

a 36.83cm (14in) height at the entrance to the ramp. The top is hinged on one side for easy lifting and 

cleaning. The ramp floor is made of wire mesh and the chamber floor should have drainage holes. 

L. maculicollis: Spotted-necked females should be separated from exhibit mates prior to parturition at 

roughly day 55 of a 60-63 day pregnancy (Schollhamer 1987), or when the female begins to show a 

tendency to keep the male or other group members away from her denning area (R.Willison, personal 

communication). Generally, females give birth to one pup, sometimes twins (Schollhamer 1987; 

R.Willison, personal communication). 

Secured sleeping dens (R.Willison, personal communication) or nest boxes that are 27" long x 20" 

wide x 15" high (68.6cm x 51cm x 38cm) have been successful (Schollhamer 1987). Typically, females do 

not use any nesting material, however, bedding should be offered to all females. 

Females should be given rubber tubs in which to swim just prior to parturition (1-3 days), and for the 

first two months or so of the pup’s life. The female will begin to bring the pup(s) out of the denning box 

when it is roughly 3-4 weeks old; at this point, she should start teaching pups to swim by placing them in 

the water tub for a few minutes at a time. It is important that the water level in tubs be kept high (which 

allows pups to hang on the lip), or that tubs/pools have a slopping ingress and egress so pups can get out 

of the water. While females are typically very vigilant, pups have suffered from hypothermia from being 

left too long in the water (Schollhamer 1987). 

P. brasiliensis: Giant otters (especially mothers) are susceptible to any human disturbance, especially 

within the surroundings of the natal den, and to discomfort created by enclosure variables (see in Sykes- 

Gatz 2005; 2.1 & 6.7; Sykes-Gatz & Gatz 2007). Several steps have been recommended to increase the 

comfort of the reproductive pair and older siblings, and improve pup-rearing success (Sykes-Gatz 2005): 

• Build a positive keeper-animal relationship and allow only familiar staff to work with the otters 

after parturition 

• The provision of food and clean water should be accomplished with minimal disturbance to the 

otters during pup rearing. 

• Cleaning should be minimal and not disruptive to the otters. 

• Reduce stress as far as possible, including loud sounds, unfamiliar people, and the introduction 

of anything new to the exhibit. 

• Prohibit visitor and zoo staff (other than immediate caretakers) access to the enclosure area. 

• Provide multiple nest box choices located in separate locations to allow parental choice according 

to their comfort level. 

• Provide appropriate substrate and exhibit conditions to include the recommended land/water 

ratio, substrate depth, digging opportunities, and dry substrate conditions. 

• Isolation of the natal den and limitation of all human activity in the vicinity at and prior to the birth 

is very important. 

• Monitoring of the natal den and early pup rearing should be done from a hidden location or 

carried out via audio and video monitoring equipment with infrared capability. 

Exhibits should be provided with multiple den sites; these can include both natural (e.g., dug by the 

otters) and man-made dens. Dens are often 48-102ft 2 (4.5-9.5m 2 ) in size. Ideally, pupping boxes just 

large enough to hold the adults, older siblings, and pups should be placed within the dens to allow the 

parents choices and maximum privacy. Dens should be provided in locations where the animals are



removed from all disturbances (Sykes-Gatz 2005). Nest box temperatures should stay above 20°C 

(68°F). Den area temperatures (where nest boxes were located) were increased to 22-23°C (71.6-73.4°F) 

during pup-rearing at one institution (Flügger 1997). P. brasiliensis in ex-situ populations have been 

observed to have a low heat tolerance (Carter & Rosas 1997; S.Sykes-Gatz & V.Gatz, personal 

observation), and pups can be very susceptible to overheating or becoming too cold. Very young pups 

especially do not thermo regulate well (Read & Meier 1996). Pups 5 months of age should not be exposed to air 

temperatures below 10°C (50°F). Parents should be prevented from taking pups outside if temperatures 

fall below these parameters. Precise recommendations for enclosure and den design are provided by 

Sykes-Gatz (2005). This publication is available on the Otter Specialist Group website 

(www.otterspecialistgroup.org) and going to the Otters in Captivity Task Force under Library. 

In at least the first days after parturition, dams have been seen to be a little protective of the pups 

when the sire tries to become involved with them. This is not abnormal behavior. Soon afterwards, the 

sire will become equally involved (and his involvement will be accepted by his mate) in the care of the 

pups. Under normal situations, it may appear that both parents and older siblings sometimes treat their 

pups a little roughly. This kind of behavior may be carried out whether otters are in the nestbox, on the 

land, or in the water. This is especially evident when parents or older siblings are teaching pups to swim. 

This seems to be normal behavior for giant otters. However, the situation should be closely monitored to 

ensure that parents and older siblings are not actually too rough with their pups, as this is abnormal. 

Starting at 2-3 weeks of age, parents will push pups under the water then let them go to resurface on their 

own or with help. This may be repeated several times to teach them to submerge (Autuori & Deutsch 

1977). Parents and older siblings have been known to teach ex-situ population-born pups to submerge 

starting at 2-6 weeks of age by holding the pup with their front feet and rolling over sideways 360° a few 

times; this has been called ‘Eskimo rolling’ (Sykes-Gatz 1999-2006). Pups also may be gently pushed or 

pulled into the water to encourage swimming. 

The following is a list of parental and older sibling behaviors observed ex-situ that often resulted in 

pup death and may be a result of sub-optimal environmental or rearing conditions: 

• Pups handled, carried, or moved to pools or new nest boxes very frequently. Generally, pups < 2 

weeks of age should not be taken into pools, if this occurs it is rare. Pups 2 weeks old or older 

should not be taken into pools more than 1-2 times per day. Older pups may tolerate more 

frequent moving, generally no more than 3 times a day. In general, pups should not be moved to 

new nest boxes on a daily basis or at most more than once or twice a day. Frequent movement of 

pups should be closely monitored without disturbance to the parents. 

• Too frequent entering of the nest box by the parents, e.g., 1-3 times per hour is normal, more can 

be abnormal. 

• Excessively forceful pushing or throwing of the pups into pools or elsewhere may be indicative of 

a problem and should be monitored, again without disturbance to the parents. In general, 

excessively forceful, rapid, or uncoordinated interactions with pups are abnormal. 

• Inappropriate mothering behavior by the dam. This may include: neglecting the pups, not lying 

still or lying incorrectly preventing pups from nursing, not staying long enough to allow for 

sufficient nursing by the pups, not visiting the pups frequently enough to allow for sufficient 

nursing, and pulling pups off their teats. These behaviors may indicate an inexperienced or 

stressed mother, or problem with lactation, such as insufficient milk production. This failure to 

produce sufficient milk amounts has been known to occur for varying periods of time as a reaction 

to stress. 

• Biting, hitting, or laying on the pups; attempted drowning of, or eating pups 

Further information on these abnormal occurrences can be obtained from Wünnemann (1995a,b), 

Flügger (1997, Autuori & Deutsch (1977), Corredor & Muñoz (2004), Sykes-Gatz (2005) and Sykes-Gatz 

and Gatz (2007). 

7.2 Artificial Insemination 

The practical use of artificial insemination (AI) with animals was developed during the early 1900s to 

replicate desirable livestock characteristics to more progeny. Over the last decade or so, AZA-accredited 

zoos and aquariums have begun using AI processes more often with many of the animals residing in their 

care. AZA Studbooks are designed to help manage otter populations by providing detailed genetic and 

demographic analyses to promote genetic diversity with breeding pair decisions within and between our



institutions. While these decisions are based upon sound biological reasoning, the efforts needed to 

ensure that transports and introductions are done properly to facilitate breeding between the animals are 

often quite complex, exhaustive, and expensive. Also, conception is not guaranteed. 

At this time AI is not used in any otter species but semen collection techniques and preservation are 

being researched by Bateman et al. (2005, 2009). 

7.3 Pregnancy and Parturition 

It is extremely important to understand the physiological and behavioral changes that occur 

throughout an otter’s pregnancy. This information is contained in Section 7.1. 

7.4 Birthing Facilities 

As parturition approaches, animal care staff should ensure that the mother is comfortable in the area 

where the birth will take place, and that this area is “baby-proofed.” This information is contained in 

Section 7.1. 

7.5 Assisted Rearing 

Although mothers may successfully give birth, there are times when they are not able to properly care 

for their offspring, both in the wild and in ex-situ populations. Fortunately, animal care staffs in AZAaccredited 

institutions are able to assist with the rearing of these offspring if necessary. 

Hand-rearing may be necessary for a variety of reasons: rejection by the parents, ill health of the 

mother, or weakness of the offspring. Careful consideration should be given as hand-rearing requires a 

great deal of time and commitment (Muir 2003). Before the decision to hand-rear is made, the potential 

for undesirable behavioral problems in a hand-reared adult should be carefully weighed (e.g., excessive 

aggression towards humans (rare in most otters), inappropriate species-specific behavior, etc.) and plans 

made to minimize deleterious effects on the development of natural behaviors as far as possible. This 

may require extensive time commitment on the part of staff, plans for fostering, relocation of the young to 

another facility, exposure to species-specific sounds, etc. At this time, the AZA Otter SSP is 

recommending hand-rearing of all otter species, if necessary. 

Pups that have been abandoned by their mother should be removed as soon as possible to prevent 

infanticide. See Chapter 6, section 6.5 for a ‘Neonatal Examination and Monitoring Protocol’. Offspring 

that are not receiving milk will be restless, possibly calling continuously, may be hypothermic, and 

scattered around the enclosure. Another indicator of trouble would be the female moving around the 

exhibit continuously while carrying the young; this could mean she is not comfortable with the denning 

provided, or there is a problem with her or the pups (Muir 2003). If it is necessary to remove offspring 

because of an exceptionally large litter, it is best to remove two of the largest pups. The temptation is 

often to take the smallest, but they stand the best chance if raised by their mother. Hand-rearing of 

singletons is more likely to lead to severe imprinting on humans than if they have a conspecific to play 

with (Muir 2003). The AZA Otter SSP recommends that singleton pups being hand-reared be placed 

together, if at all possible. To date, fostering has been attempted once with otter pups and was 

successful. A pup was taken from a female with no milk and sent to another facility where their female 

was already nursing pups. In these cases, the AZA Otter SSP management team should be consulted 

first. Other institutions have been successful at supplement feeding pups left with their mother. Young 

otters removed for hand-rearing should not routinely be reintroduced to the parents with an expectation of 

acceptance. Introductions of hand-reared animals should follow procedures specified in the 

Introduction/Reintroduction section. 

Physical Care Protocol: Incubators provide the best source of warmth. Heat lamps are too intense and 

can be dehydrating. In an emergency, hot water bottles wrapped in a towel may be placed in a box with 

the pups nestled next to it, or they can be warmed slowly by placing them next to your body (Muir 2003). 

Pups may feel more secure if wrapped in layers of towels; this also aids in keeping them warm (Muir 

2003). Pups should be dried after feeding/bathing to prevent hypothermia until they are proficient at selfgrooming. 

The normal body temperature for pups is unknown, but the animal should feel warm to the 

touch. 

Altricial young are unable to self-regulate their body temperature during the early postnatal period and 

require an external source of warmth. If an incubator is not used, it may be necessary to place a heating 

pad, set on low, under the housing container until the pups are able to thermo regulate. Meier (1986) and 

Wallach & Boever (1983) recommend 29.4-32.2°C (85-90°F) and 50-60% humidity as the desired



incubator setting for neonate mustelids. The temperature should be gradually reduced to room 

temperature, 21.2-23.9°C (70-75°F), over the course of about three weeks (unless the neonate becomes 

ill). Litters of pups are less likely to need additional ambient heat since huddling together may provide an 

adequate amount of warmth. External temperatures should be closely monitored to prevent hyperthermia. 

Rapid and/or open-mouth breathing, restlessness, and hair loss are indication of an external environment 

that is too warm. 

Pups should be stimulated to urinate and defecate at least 4-5 times each day for several weeks, 

generally before feeding. However, some animals may respond better to post-feeding stimulation. The 

genitals and anal area are rubbed gently with a finger, towel, or damp cotton to stimulate the baby to 

urinate and have a bowel movement. If pups do not urinate and/or defecate after two successive 

feedings, the formula should be reviewed and their health status evaluated immediately. 

Specific environmental parameters, formula information, etc. for hand-rearing L. canadensis and P. 

brasiliensis pups can be found in the North American River Otter Husbandry Notebook, 2 nd Edition (Reed- 

Smith 2001) and International Giant Otter Studbook Husbandry and Management Information and 

Guidelines (Sykes-Gatz 2005), respectively; these are available on the Otter Specialist Group web site 

(www.otterspecialistgroup.org). The hand-rearing of giant otters (P. brasiliensis) is somewhat different 

than that of other otter species, because their development is slower. Detailed information on the types of 

records needed, signs of illness, etc. are available in the Giant Otter Husbandry Manual (Sykes-Gatz 

2005). 

Feeding Amount and Frequency: Initially, the animal should receive only an electrolyte solution for the 

first 2-3 feedings, depending on how compromised it is. This is to rehydrate the animal and clear the 

stomach of the maternal milk. The artificial formula should be started at a diluted concentration, generally 

at a 1:4 ratio (mixed formula: water) for another 2-3 feedings. It generally takes about 72 hours to get the 

animal on full-strength formula by gradually offering higher concentrations. Depending on the species, 4-5 

feedings of each concentration level (1:2, 1:1, 2:1, full-strength) are required to allow for adaptation and to 

minimize the onset of digestive problems, particularly diarrhea. During the initial phase (24-36 hours), 

weight loss is to be expected, but the animal should quickly begin to maintain weight and then start 

gaining as the formula concentration increases. It is important that the pups are not given full strength 

formula too soon (in less than 48 hours after pulling for hand-rearing) because the likelihood of diarrhea is 

extremely high. Diarrhea is of particular concern with neonates less than one week of age, because they 

have very little or no immunity to infections. 

Pups should have a normal body temperature and be properly hydrated before starting them on 

formula. Young mammals require a specific amount of calories per day for optimum development and 

growth. A nutritionally dense milk formula will allow for fewer feedings than more dilute formulas that are 

low in fat or protein. A method for calculating the volume of food to be offered per meal as well as total 

daily amount is presented below. 

The Basal Metabolic Rate (BMR) or Basal Energy Requirement (BER) is the amount of energy (kcal) 

an animal needs for basic metabolic function at rest in a thermo-neutral zone. This represents the amount 

of calories it needs to stay alive, without having to use energy to maintain normal body temperatures 

(Grant 2004). Mustelids have a higher metabolic rate per body weight than many other placental 

mammals. For that reason, Iversen’s equation of 84.6 x body weight (in kg) 0.78 (Iversen 1972) is used 

rather than Kleiber’s equation of 70 x body weight (in kg) 0.75 (Kleiber 1947) typically used for other 

species. Therefore, for a 200g river otter, the BER would be: 84.6 x 0.2 0.78 = ~24 kcal/day. 

Once the BER is established, the Maintenance Energy Requirement (MER) can be calculated. This 

measurement determines the amount of calories the animal needs to function in a normal capacity at its 

life stage. For adults in a maintenance life stage, the BER is multiplied by 2. For pups that have a higher 

metabolism and are developing and growing, the BER is multiplied by 3 or 4 (Evans 1985), depending on 

the species and other factors. 

The stomach capacity for most placental mammals is 5-7% of the total body weight (Meehan 1994). 

Convert the body weight into grams to find the stomach volume in ml (cc). To calculate the stomach 

capacity in ounces, convert body weight into grams (30g ~ 1 oz). It is important that units are the same for 

body weight and stomach volume. The stomach capacity is the amount of formula an infant can 

comfortably consume at one feeding. Offering much more than this value may lead to overfilling, stomach 

distension, and bloat. It also prevents complete emptying of the stomach before the next feeding, and 

promotes the overgrowth of potentially pathogenic bacteria, diarrhea, and enteritis (Evans 1985).



The following calculations will determine the total volume and kcal to feed/day, as well as the amount 

of formula for each feeding and the total number of feedings daily. 

• Calculate Maintenance Energy Requirement: 84.6 x body wt (kg) 0.78 x 3. 

• Determine stomach capacity (amount that can be fed at each meal): Body weight (in grams) x 

0.05. 

• Divide Maintenance Energy Requirement (number of calories required per day) by the number of 

kcal/ml in the formula to determine the volume to be consumed per day (this can be converted 

into ounces by dividing it by 30). 

• Divide ml of formula per day by volume to be consumed at each meal (stomach capacity). This 

gives the number of meals to offer per day. 

• Divide 24 hours by the number of feedings/day to find the time interval between feedings. 

• See Table 12. 

Table 12: Calculations for formula volume and feeding frequency for neonate with an approximate birth weight of 

135g (MER = Maintenance Energy Requirement) 

Step 1: calculate MER 84.6 x 0.135kg 0 .78 x 3 ~53 kcal/day 

Step 2: determine stomach capacity 135g x 0.05 (stomach capacity of 

5% body weight) 

Step 3: calculate daily volume fed 53 kcal/day (MER) 

1.78 kcal/ml (formula contents) 

Step 4: number of feedings 30ml/day (total volume fed) 

7ml/feeding (stomach capacity) 

~7g (ml) per feeding 

~30ml/day 

4.2 feedings/day (=5) 

Step 5: feeding schedule 24 hrs/5 feedings Every 5 hours 

New calculations should be performed every few days so formula volume can be adjusted to 

accommodate growth. The general target average daily gain for infants is 5-8% increase of body wt/day 

while on formula feeding and 8-10% body wt increase/day on weaning diet (Grant 2005). Since neonates 

being hand-reared (less than one week of age) are typically severely compromised, they should be given 

smaller, more frequent feedings than calculated until roughly 2-4 weeks of age. 

As a general rule, animals should have an overnight break between feedings that are no longer than 

twice the time period between daytime feedings (equivalent to missing one feeding). For example, if they 

are being fed every three hours during the day, they can go six hours at night without food. When they 

are eating every four hours, they can go eight hours at night. It is not advisable to go more than eight 

hours between feedings with species that typically nurse throughout the day when mother-raised. 

Intervals between feeding also will depend on how healthy or strong the infants are. Very weak neonates 

will probably need feedings every few hours even through the night; typically this is necessary for only a 

few days to a week. The AZA Otter SSP recommends that neonates be fed every two hours around the 

clock initially. Depending on how the animal is doing, these feedings may be stretched to every three 

hours after the first few weeks. 

Otter pups should only be fed if the pup is hungry and suckling vigorously. Weak infants may be 

hypothermic, dehydrated and/or hypoglycemic. Do not offer anything by mouth until the body temperature 

is within the normal range for its age (i.e., warm, not hot, to the touch). Electrolytes can be offered orally if 

the pup is suckling, or subcutaneously if it is too weak; 2.5-5% dextrose can also be given to raise the 

pup’s glucose level. More research is required to determine body temperature norms for young of all the 

otter species. This information should be collected by all facilities hand-rearing otter pups and submitted 

to the AZA Otter SSP. Young animals will be hungry at some feedings, less at others, but this is quite 

normal (Muir 2003). However, refusal of two feedings is a sign of trouble in young otters. Pups will not die 

from being slightly underfed, but overfeeding may result in gastrointestinal disease, which is potentially 

fatal. 

If any animal's formula is changed abruptly, it is likely to cause diarrhea, which can dehydrate the pup 

quickly. Any formula changes should be made slowly, by combining the formulas and gradually changing 

the ratio from more of the first to more of the second. If an animal develops diarrhea or becomes 

constipated with no change having been made in the formula, consult the veterinarian. In general, 

adjusting the formula ratios should be attempted before medicating the animal. For diarrhea, increase the



ratio of water to all the other ingredients. Be sure the water has been boiled or sterilized well, and the 

bottle is clean. Subcutaneous fluids (e.g., lactated ringers) may be needed if the infant dehydrates 

significantly. 

Feeding Techniques: To bottle feed, hold the pup in the correct nursing position; sternally recumbent 

(abdomen down, not on its back), with the head up. Place the hand holding the bottle in such a way that it 

provides a surface for the pup to push against with its front feet. If milk comes through their nose, the 

nipple hole may be too large or the pup may be trying to eat too quickly. Make sure there is consistency 

with who is feeding the pups. Note any changes in feeding immediately. Decreased appetite, chewing on 

the nipple instead of sucking, or gulping food down too quickly can be signs of a problem (Blum 2004). 

It is important to keep in mind that neonates are obligate nose breathers and incapable of breathing 

through their mouths and nursing at the same time. For this reason, respiratory infections can be life 

threatening because they may interfere with breathing and make nursing difficult or impossible (Meier 

1985). Aspirated formula is frequently a contributing factor to neonatal respiratory infections; to avoid this, 

be sure to select the appropriate nipple. The nipple’s hole needs to suit the neonate’s sucking reflex. 

Also, if a nipple is too stiff, the pup may tire and refuse to nurse. 

If an animal aspirates fluids the recommended protocol is to hold the infant with head and chest lower 

than the hind end. A rubber bulb syringe should be used to suck out as much fluid from the nostrils and 

the back of the throat as possible. If aspiration is suspected, or if fluid is heard in the lungs, contact the 

veterinarian immediately; do not administer drugs without the veterinarian’s involvement. Monitor body 

temperature closely for the occurrence of a fever and a decline in the animal’s appetite and general 

attitude. Depending on the condition and age of the animal, diagnostic procedures may include 

radiographs, CBC, and chemistry. It is possible to start a course of antibiotics while results from the 

bloodwork are pending, and the attending veterinarian can prescribe an appropriate antibiotic course. 

Pups will need to be stimulated to urinate and defecate for the first six weeks of life, either 

immediately before or after feeding. Parent-reared giant otters of ex-situ populations are reported to have 

required stimulation by their parents to urinate or defecate for up to 10 weeks of age. In at least one case, 

a hand-reared individual needed to be stimulated to urinate/defecate until it was 2.5-3 months old (Sykes- 

Gatz 1999-2006). In other cases (Corredor & Muñoz 2004), pups were reported using latrines on their 

own at 9 weeks of age. Some pups also may require “burping” to prevent gas build-up in the abdomen. 

P. brasiliensis: One of the most reliable methods of determining if the young are nursing successfully is 

monitoring for what Sykes-Gatz (2005) calls the “nursing hum”, which pups make when they are suckling. 

This hum is a somewhat higher pitched and faster vocalization than the contact hum described by Duplaix 

(1980), which has a twittering quality to it. The nursing hum is performed when a pup is nursing from the 

mother or a bottle. Sykes-Gatz (2005) also reports this call, when given by a caregiver, can encourage 

pups to feed. From birth, the pups also display “tail wagging” when they nurse, wagging their tails rather 

quickly and repeatedly from side-to-side. Some individuals may require “burping” to prevent gas build-up 

in the abdomen. For more detailed information refer to Sykes-Gatz (2005). 

A. cinereus: Pups may be slow to learn how to suckle from a bottle, in one case taking eight days before 

suckling without aspirating (Webb 2008). Care should be taken to ensure that the nipple’s hole is not too 

large and that pups are fed slowly at the beginning. For additional information see Webb 2008 (also 

available online at (www.otterspecialistgroup.org/Library/TaskForces/OCT.html). 

Hand-rearing Formulas: It is important that the artificial milk formula matches the maternal milk in 

protein, fat, and carbohydrate composition as closely as possible. Table 13 provides information on the 

nutritional content of otter milk, and Table 14 provides information on the nutritional composition of 

selected substitute milk formulas/replacers. 

Table 13: Otter (Lutra spp.) Milk Nutrition Composition on As Fed (AFB) and Dry Matter Basis (DMB) (Ben Shaul 

1962; Jenness & Sloan 1970) 

Species Solids % Kcal (ml) Fat % Protein % Carb. % 

Otter 38.0 2.6 (AFB) 

24.0 (AFB) 

63.2(DMB) 

11.0 (AFB) 

28.9(DMB) 

0.1 (AFB) 

0.3 (DMB) 

Esbilac ® (or Milk-Matrix ® 33/40) is preferred as the base for milk formulas offered to otters and 

provides good pup growth. The addition of Multi-Milk ® (or Milk-Matrix ® 30/55) increases the total fat and



protein content without adding substantially to the carbohydrate content of the formula. The maternal milk 

composition of otter milk only has a trace amount of milk sugars, so this component of the substitute 

formula should be kept as low as possible to prevent gastric upset and diarrhea. See Table 14 on the 

following next page.

Table 14: Nutritional analysis of commercial animal milk replacers 

Product 

Solids 

% 

Fat 

% 

Protein 

% 

Carbohydrates 

% 

Ash 

% 



Energy 

(KCAL/ML) 

Esbilac 

Undiluted powder 95.00 40.00 33.00 15.80 6.00 6.20 

Diluted 1:3* 15.00 6.00 4.95 2.38 0.90 0.93 

Diluted 1:1.5* 30.00 12.00 9.90 4.76 1.80 1.86 

Liquid product 15.00 6.00 4.95 2.38 0.90 0.93 

KMR 

Undiluted powder 95.00 25.00 42.00 26.00 7.00 5.77 

Diluted 1:3* 18.00 4.50 7.56 4.68 1.26 1.04 

Diluted 1:1.5* 36.00 9.00 15.12 9.36 2.52 2.07 

Liquid product 18.00 4.50 7.56 4.68 1.26 1.04 

Multi-Milk 

Undiluted powder 97.50 53.00 34.50 

0 

6.63 6.85 

Diluted 1:1* 22.70 12.00 7.83 0 1.51 1.55 

Diluted1.5:1* 36.00 19.59 12.75 0 2.54 2.47 

Evaporated Milk 

Undiluted product 

22.00 

7.00 

7.90 

Multi-Milk:KMR+ 

1:1* 

22.81 8.93 8.71 

3.20 

1.55 1.45 

3:1* 22.90 10.97 8.63 1.54 1.59 1.57 

4:1* 22.90 10.90 8.27 1.17 1.50 1.51 

1:3* 22.70 7.28 9.10 4.39 2.30 1.37 

1:4* 22.60 6.95 9.16 4.68 1.57 1.36 

Multi-Milk:KMR++ 

1:1* 

34.22 13.40 13.07 

4.80 

2.33 2.18 

3:1* 34.55 16.46 13.03 2.31 2.39 2.36 

4:1* 34.55 16.35 12.41 1.76 2.25 2.28 

1:3* 34.05 10.92 13.65 6.59 3.45 2.06 

1:4* 33.90 10.43 13.74 7.02 2.36 2.04 

Multi-Milk:Esbilac+ 

1:1* 

22.81 10.63 7.70 

1.78 

1.44 1.49 

3:1* 22.93 11.63 8.00 0.89 1.52 1.56 

4:1* 22.90 11.60 7.86 0.71 1.49 1.55 

1:3* 22.70 9.81 8.75 2.67 2.13 1.51 

1:4* 22.60 9.65 7.54 2.84 1.39 1.43 

Multi-Milk:Esbilac++ 

1:1* 

34.22 15.95 11.55 

2.67 

2.16 2.24 

3:1* 34.40 17.45 12.00 1.34 2.28 2.33 

4:1* 34.35 17.40 11.79 1.07 2.24 2.33 

1:3* 34.05 14.72 13.13 4.01 3.20 2.28 

1:4* 33.90 14.48 11.31 4.26 2.09 2.15 

* Ratio of powder to water; + Ratio of powder-to-powder, diluted 1 part powder to 1 part water; ++ Ratio of powder-to-powder, 

diluted 1.5 parts powder to 1 part water (Evans 1985) 

The addition of an anti-gas build-up product to the formula should be considered (milk sugars can 

cause the build-up of gas). Lact-aid ® is an enzyme that has been used successfully with many species. 

Add two drops of Lact-aid ® to 100ml of mixed formula. The formula then should be refrigerated for 24 

hours prior to feeding for the enzyme to perform correctly (Grant 2005). Lactobacillus spp., in Bene-bac ® 

or Probios ® , is a group of beneficial gut bacteria that also break down milk sugars in the digestive tract. 

Follow label instructions for these products. 

9.70 

0.70 

1.49



Table 15: Substitute milk formulas for otters. Values taken from product composition documents available from 

PetAg (K.Grant, personal communication) 

% % % 

Formula 

% Carb Kcal/ml 

Solids Fat Protein 

Formula #1 

1 part Esbilac ® or Milk Matrix ® 33/40 

1 part Multi-Milk ® or Milk Matrix ® 30/55 

2 parts water 

Formula #2 

1 part Multi-Milk ® or Milk Matrix 30/55 ® 

1 part water 

30.9 15.6 10.5 2.7 1.78 

31.3 17.8 10.4 1.1 1.91 

L. Canadensis: At this time, the preferred formula is canned Esbilac ® due to palatability and good pup 

growth. Milk Matrix ® based formulas also are nutritionally suitable but some facilities have had pups 

refuse this formula (Blum 2004) while others have had good success. 

Weaning: Some of the following recommendations do not apply to P. brasiliensis (e.g. offering food in a 

bowl). The weaning process should be started when the pup shows interest in solid food, generally at 

about eight weeks of age. If the pup is not gaining enough weight on formula alone, solid food can be 

added at six weeks of age (this may need to be pureed or chopped). To begin, formula can be mixed with 

AD diet (canned cat food or similar), baby food, mashed up fish, rice cereal, or ground meat. New food 

can be added to the bottle; feed this mixture with a syringe, baby bottle, or offer it in a bowl. Do not 

provide milk formula in a bowl to giant otters, as they tend to inhale liquids into the nose until they 

become proficient at eating solid foods (McTurk & Spelman 2005). Only add one new food component to 

their diet every couple of days until they are eating solids well. It is best to be creative, flexible, and not to 

rush the weaning process. In the case of problems, try different approaches, try them multiple times, and 

try foods in new ways like bottles, syringes, suction bulbs, bowls, etc. Do not cut back on bottle-feeding to 

make the pup “hungry”. Offer new food at the beginning of the feeding and finish with the bottle (Blum 

2004). Situations to watch for during the weaning process include (Blum 2004): weight loss, diarrhea and 

sucking behavior. If sucking on tails, feet, genitals, etc. is observed between feedings, an additional 

bottle-feeding should be offered for a few days. R. Green of the Vincent Wildlife Trust recommends 

putting orange oil on the genitals to discourage sucking; this worked well with Lutra lutra and is not 

harmful to the otter (G.Yoxon, personal communication). 

Swimming, Terrestrial Activities, and Behavioral Stimulation: Otter pups are not born knowing how to 

swim and may even be scared of the water. They will usually start to take interest in the water at 4-8 

weeks of age. The pups should be started off in shallow pools and watched carefully; once comfortable, 

they can gradually be introduced to deeper water. Pups should be dried off completely and warmed after 

their swim. 

Enrichment is crucial to the development of the pups; toddler safe toys, grooming materials, dens, 

climbing structures, live food, etc. have all been used successfully. The more items they are introduced to 

otters at an early age, the more they will interact with as they age. All toys need to be safe and approved 

by the veterinary staff. The suitability of toys should be regularly re-evaluated, as some may no longer be 

safe as the otter grows. Due to the tendency of all otters to take things into the water, the use of 

cardboard or other paper-type items, especially for young animals, is not recommended. Cases of these 

items becoming water logged and congealing in an animal’s mouth or over their nose have been 

reported. 

Pup Development: The following information provides a summary of pup development. More specific 

information can be found in the Otter Husbandry Manuals (Lombardi et al. 1998; Reed-Smith 2001; 

Sykes-Gatz 2005). See Appendix I for pup weight charts. 

A. cinereus: 

• Eyes begin opening at between 17 and 28 days, fully open by day 45 

• Teeth begin erupting about day 20 and canines erupting ~ day 91 (Webb 2008) 

• Thermo regulating well on their own about day 38 (Webb 2008) 

• Moving on their own between day 39 and day 50



• Urinating and defecating on their own (hand reared animals) by day 59 (Webb 2008) 

• Generally born with mostly grayish fur, darkens by 6-7weeks 

• Solid food 7-8 weeks; weaned 82-120 days 

• Hand reared animals eating solid well by day 92 and weaned on day 130 at a weight of 2336 

grams (Webb 2008) 

A. capensis: At this time there is no information available on pup development. More research is required. 

L. maculicollis: Pups are born with white on their lips. After a few days, patches of white/orangish colored 

hair develop on their chest or groin area. These patches change to an orange color, before changing 

back to cream or white as the pups reach full growth or maturity. The age at which these color changes 

occur appears to be highly variable and is currently being documented (D.Benza, personal 

communication; R.Willison, personal communication). 

• First spots seen ~6 days, whitish but turned orange in a few days. More orange spots developed 

by day 42 

• Eyes open at 34-46 days 

• First crawling at about 20 days, crawling well 42 days 

• First teeth erupting at 23-29 days, all teeth in ~78 days 

• Walking well at ~ 37 days, running 59 days 

• Leaving den on own at about 57 days 

• Playing in water bowl ~ 61 days 

• First going in to water on their own at about 57-91 days; variation comes from water tub versus 

pool exploration 

• First pool swimming lessons ~ 86 days (timing may be due to when family is allowed into the 

exhibit) 

• First eat solids at about 60-73 days 

L. canadensis: Consult the North American River Otter Husbandry Notebook 1 st & 2 nd editions for more 

detailed information (Reed-Smith 1994, 2001). 

• Birth weight: 120-135g 

• Born blind with dark brown fur 

• External ears are flat against the head, and claws and toe webbing are well formed. 

• Deciduous upper and lower canines erupt at about 12 days 

• Eyes fully open at 28-35 days 

• Walking at about 35-42 days, first swimming lesson generally at 28-56 days 

• Beginning to play ~25-42 days 

• Leaving nest box on their own ~49 days 

• Pelt change 28-56 days, born with all dark fur 

• First solid food taken at 42-56 days 

• Localized latrine use ~49 days 

• Pups should be weaned by 3-4 months of age 

P. brasiliensis: Because this species requires complete isolation and privacy (particularly primiparous 

pairs), detailed information on pup development is taken from video and audio recordings. Sykes-Gatz 

(2005) provides more detail on pup rearing and development. McTurk & Spelman (2005) also provide 

information on hand-rearing and rehabilitation of orphaned giant otters. An outline of giant otter pup 

development is provided below (Wünnemann 1990, 1995a,b; McTurk & Spelman 2005; Sykes-Gatz 2005, 

1999-2006; V.Gatz, personal communication; N.Duplaix, personal communication): 

• Weight at birth – 150-265g 

• Birth pelt is grayish in color and darkens by 6-7 weeks of age 

• Eyes begin opening at ~28 days and are fully open by ~45 days 

• Pups should be moving on their own by 39-50 days 

• First leave the nest box on their own at 63-67 days 

• First swimming lessons at 20-60 days, or as early as 11 days 

• Pups can be reliably sexed at 10 weeks 

• Pups swim on their own for the first time at 63-67 days



• Pups will begin playing with solid food at roughly 56 days, but generally do not consume any until 

about 70-90 days. 

• Pups will begin weaning at roughly 4 months of age, but can nurse insignificant amounts (this 

provides little nutritional value) at 6.5 to 8 months of age. 

• Fish should first be offered pups at 2.5-4 months of age 

• 100% of their required caloric intake should be offered in formula/mother’s milk form until roughly 

2.5 months of age 

• Pups should be weaned from formula between 6.5-10 months of age 

• Pups should be weaned on a fish based diet; rice cereal has been used successfully as a dietary 

addition for hand-reared pups. Formula should not be offered in a bowl, as giant otters tend to 

inhale liquids into the nose until they are proficient at eating solid foods. 

• Pups are approximately ¾ the size of adults at 10 months of age, although this will vary 

7.6 Contraception 

Many animals cared for in AZA-accredited institutions breed so successfully that contraception 

techniques are implemented to ensure that the population remains at a healthy size. In addition to 

reversible contraception, reproduction can be prevented by separating the sexes or by permanent 

sterilization. In general, reversible contraception is preferable because it allows natural social groups to 

be maintained while managing the genetic health of the population. Permanent sterilization may be 

considered for individuals that are genetically well-represented or for whom reproduction would pose 

health risks. The contraceptive methods most suitable for otters are outlined below. More details on 

products, application, and ordering information can be found on the Intstitution E webpage: 

www.stlzoo.org/contraception. 

The progestin-based melengestrol acetate (MGA) implant, previously the most widely used 

contraceptive in zoos, has been associated with uterine and mammary pathology in felids and suspected 

in other carnivore species (Munson 2006). Other progestins (e.g., Depo-Provera ® , Ovaban ® ) are likely to 

have the same deleterious effects. C.Osmann (personal communication) specifically recommends against 

using progestins in P. brasiliensis for the reasons mentioned above and because these side effects may 

compromise future breedings. For carnivores, one institution now recommends GnRH agonists, e.g., 

Suprelorin ® (deslorelin) implants or Lupron Depot ® (leuprolide acetate), as safer alternatives. Although 

GnRH agonists appear safe and effective, dosages and duration of efficacy have not been systematically 

evaluated for all species. GnRH agonists can be used in either females or males, and side effects are 

generally those associated with gonadectomy, especially weight gain, which should be managed through 

diet. Suprelorin ® was developed for domestic dogs and has been used successfully in African clawless 

otters, North American river otters, Asian small clawed otters and sea otters. 

Gonadotropin Releasing Hormone (GnRH) Agonists: GnRH agonists (e.g., Suprelorin ® implants or 

Lupron Depot ® ) achieve contraception by reversibly suppressing the reproductive endocrine system and 

preventing production of pituitary (FSH and LH) and gonadal hormones (estradiol and progesterone in 

females and testosterone in males). The observed effects are similar to those following either ovariectomy 

in females or castration in males, but are reversible. GnRH agonists first stimulate the reproductive 

system, which can result in estrus and ovulation in females or temporary enhancement of testosterone 

and semen production in males. Then, down-regulation follows the initial stimulation. The stimulatory 

phase can be prevented in females by daily Ovaban administration for one week before and one week 

after implant placement (Wright et al. 2001). 

GnRH agonists should not be used during pregnancy, since they may cause spontaneous abortion or 

prevent mammary development necessary for lactation. They may prevent initiation of lactation by 

inhibiting progesterone secretion, but effects on established lactation are less likely. New data from 

domestic cats have shown no effect on subsequent reproduction when treatment began before puberty; 

no research in prepubertal otters has been conducted. 

A drawback of these products is that time of reversal cannot be controlled. Neither the implant 

(Suprelorin ® ) nor the depot vehicle (Lupron ® ) can be removed to shorten the duration of efficacy to time 

reversals. The most widely used formulations are designed to be effective for either 6 or 12 months, but 

those are for the most part minimum durations, which can be longer in some individuals. 

Although GnRH agonists can also be an effective contraceptive in males, they are more commonly 

used in females. This is because monitoring efficacy by suppression of estrous behavior or cyclic gonadal



steroids in feces is usually easier than ensuring continued absence of sperm in males, since most 

institutions cannot perform regular semen collections. Suprelorin ® has been tested primarily in domestic 

dogs, whereas Lupron Depot ® has been used primarily in humans, but should be as effective as 

Suprelorin ® since the GnRH molecule is identical in all mammalian species. 

If used in males, disappearance of sperm from the ejaculate following down-regulation of testosterone 

may take an additional 6 weeks, as with vasectomy. It should be easier to suppress the onset of 

spermatogenesis in seasonally breeding species, but that process begins at least 2 months before the 

first typical appearance of sperm. Thus, treatment should be initiated at least 2 months before the 

anticipated onset of breeding. 

Progestins: If progestins (e.g., Melengestrol acetate (MGA) implants, Depo-Provera ® injections, 

Ovaban ® pills) have to be used, they should be administered for no more than 2 years and then 

discontinued to allow for a pregnancy. Discontinuing progestin contraception and allowing non-pregnant 

cycles does not substitute for a pregnancy. Use of progestins for more than a total of 4 years is not 

recommended. MGA implants last at least 2 years, and clearance of the hormone from the system occurs 

rapidly after implant removal. Progestins are considered safe to use during lactation. 

Vaccines: The porcine zona pellucida (PZP) vaccine has not been tested in otters, but may cause 

permanent sterility in many carnivore species after only one or two treatments. This approach is not 

recommended. 

Ovariectomy or Ovariohysterectomy: Removal of ovaries is a safe and effective method to prevent 

reproduction for animals that are eligible for permanent sterilization. In general, ovariectomy is sufficient 

in young females, whereas removal of the uterus as well as ovaries is preferable in older females, due to 

the increased likelihood of uterine pathology with age. 

Vasectomy: Vasectomy of males will not prevent potential adverse effects to females that can result from 

prolonged, cyclic exposure to the endogenous progesterone associated with the pseudo-pregnancy that 

follows ovulation. This approach is not recommended for otters.



Chapter 8. Behavior Management 

8.1 Animal Training 

Classical and operant conditioning techniques have been used to train animals for over a century. 

Classical conditioning is a form of associative learning demonstrated by Ivan Pavlov. Classical 

conditioning involves the presentation of a neutral stimulus that will be conditioned (CS) along with an 

unconditioned stimulus that evokes an innate, often reflexive, response (US). If the CS and the US are 

repeatedly paired, eventually the two stimuli become associated and the otter will begin to produce a 

conditioned behavioral response to the CS. 

Operant conditioning uses the consequences of a behavior to modify the occurrence and form of that 

behavior. Reinforcement and punishment are the core tools of operant conditioning. Positive 

reinforcement occurs when a behavior is followed by a favorable stimulus to increase the frequency of 

that behavior. Negative reinforcement occurs when a behavior is followed by the removal of an aversive 

stimulus to also increase the frequency of that behavior. Positive punishment occurs when a behavior is 

followed by an aversive stimulus to decrease the frequency of that behavior. Negative punishment occurs 

when a behavior is followed by the removal of a favorable stimulus also to decrease the frequency of that 

behavior. AZA-accredited institutions are expected to utilize reinforcing conditioning techniques to 

facilitate husbandry procedures and behavioral research investigations. 

Otters are excellent candidates for behavioral training programs focusing on routine and non-routine 

husbandry tasks, such as shifting, weighing, entering squeeze cages or crates, stationing for close visual 

inspections or injections, etc. Standard positive reinforcement behavioral training techniques are used 

successfully on river otters at numerous facilities. As far as possible, all animals should routinely shift into 

a holding area and readily separate into specific holding areas on cue. Animals should be trained to come 

to the keeper when called for daily health checks, and remain calm and not aggressive during these 

checks. 

Keepers should avoid use of aversive stimuli in the daily management of otters. Profound aversive 

stimuli such as squirting with hoses, loud noises, harsh words, and long-term withholding of food are 

inappropriate unless serious injury of keeper or animal is imminent (e.g., serious fight). In general, otters 

respond to profound aversive stimuli with fear and/or aggression. It is best to maintain positive and 

pleasant keeper/animal interactions. Assessing the animal’s motivation (e.g., why should it “want” to 

come in? Why does it “want” to stay outside? What is the animal’s motivation, and how does it relate to 

the animal’s behavior in the wild?) is a useful exercise when training problems occur. Patience and 

planning are keys to success (Wooster 1998). See Table 16 for a list of commonly trained otter behaviors, 

as well as relevant cues and criteria. Successful training programs include those that involve establishing 

training goals set by the entire staff. These goals include a list of behaviors that facilitate desired 

husbandry procedures. Goals are accomplished by developing training plans that define training steps, 

cues, and criteria for the desired behaviors. Progress of training plans should be monitored and 

evaluated. Once desired behaviors are achieved, they should be maintained with practice on a regular 

daily basis. See Appendix J for additional training information. 

Otters can be trained through positive reinforcement for almost all behaviors required for husbandry 

procedures, whether it is routine or a not-so common event. Non-routine husbandry behaviors can 

include procedures such as hand injection, ultrasound, nipple manipulation/milk collection (for larger 

species in particular), and tactile body exams. A P. brasiliensis female was successfully trained to allow 

manual milk pumping and ultrasounds to detect pregnancy and uterine condition (Gatz 2002). Giant otters 

also have been trained to allow tactile body exams, the taking of body temperature, weight, heart and 

respiration rates, as well as to participate in other husbandry procedures (Sykes-Gatz 2005). 

Otters respond quickly to voice commands via operant conditioning. Training can be done on or off 

exhibit. Otters respond to a protected contact and free contact situation. In general, the otter species 

should be trained in a protected contact situation (i.e., keeper and animal should be separated by a mesh 

barrier). Exhibits should be designed with mesh at a particular area specifically for training. There are 

some species (A. cinereus) or cases (L. canadensis particularly males) where an institution feels that 

protected contact training may not be called for, but these decisions should be carefully evaluated on an 

ongoing basis. If institutional philosophy permits, otters can be a part of an educational talk or keeper talk 

in a free contact area within their exhibit.



It is recommended that all facilities have holding areas in order to shift animals into/out of their 

primary enclosure. Husbandry training may occur anywhere the individual animal seems to feel 

comfortable, and where the keeper can safely access them. Managers and caretakers should decide if 

food rewards can be hand fed or if a meat stick should be used to deliver the food. 

The following table (Table 16) provides some examples of husbandry training cues and criteria for 

behaviors trained with otters at various AZA institutions. Mckay (2009) describes some basic approaches 

to training otters. 

Table 16: Sample behaviors & training cues for otters (provided by: *Institution F; **Institution G; ***Institution H; 

^Institution I; ^^Institution J; & +Institution K). Behaviors not identified are trained at all reporting institutions. 

Behavior Verbal cue Visual cue Criteria for reinforcement 

Down 

* 

Up 

* 

Up 

^ 

Stand 

** 

Kennel 

* 

Entering a crate 

** 

Squeeze/Crate 

***/^^ 

Crate 

+ 

Scale 

* 

Target 

* 

Target 

** 

Target 

***/ ^ 

Target 

+ 

Stay 

** 

Stay/remote 

stay 

^ 

“down” Hand flat in front of abdomen- 

moved in a downward motion 

“up” Index finger moved in upward 

motion to place you want them to 

target to 

Animal lays down quietly 

Animal moves to position of index 

finger 

“up” Left index points into the air Animal stands up 

“up” Use left hand and give the thumbsup 

sign 

“in” Index finger used to point into the 

kennel 

“box” Hand begins in fist in front of chest. 

As command is said, swing arm out 

and up in direction of the box and 

open hand into a high five. 

“crate” Target into squeeze cage or point 

to crate 

Hand placed on chain link near 

back of crate 

Otter keeps both back feet on the 

ground while standing up against 

the cage. Front feet should be 

hanging onto target pole place 

against the bars. 

Animal goes in kennel and allows 

door to be closed 

Animal will enter crate and lie down 

at the far right end. Animal will wait 

in position until bridged. 

Animal enters and allows the door 

to be closed 

Animal enters and stands in the 

crate, tail completely in 

“scale” Index finger used to point to scale Animal gets on scale & waits 

“here” Closed fist presented to front of 

mesh 

Nose placed at position of fist 

“target” Hold up target pole Animal grabs with both hands 

without biting – ASC otters 

“target” Show target pole Nose placed on target and holds 

until bridged 

Show 15’ broom handle on fence Put nose to target 

“stay” Right hand palm down and out. 

While in this position, push slightly 

toward animal while saying verbal 

cue. 

“stay” Hold hand up, palm towards the 

animal 

Hold fist up 

Animal stands/sits still while trainer 

moves away and returns 

Animal stays calmly 

Hold 

^^ 

“hold” Hand cue Animal stays in place 

Lying parallel to ‘lie” Palm flat out and facing down. Animal lays down parallel to and


cage front 

Shift 

Come in 

* 

Recall 

+ 

Station 

**/^^ 

Station 

^ 

Follow 

Foot present 

Paw 

Ultrasound 

** 

Ultrasound 

*** 

Paint 

*** 

Nipple 

presentation 

*** 

Ventral present 

+ 

Jumping into the 

pool 

** 

Sweep arm in direction animal 

should face. 

“over” Arm begins up and parallel to chest, 

index finger pointed up. (Use arm 

that is in the direction you want the 

animal to shift. Move arm and 

corresponding foot in a sweeping 

motion indicating the direction you 

want the animal to go). 

3 whistles- flat 

tone 



touching cage front. Remains calm 

and quiet until bridged. 

The animal goes to the are 

indicated, comes to front of cage, 

stands quietly with eyes on trainer 

None Animal moves in to location of 

person whistling 

Clicker Animal moves off exhibit to catch 

area 

None Trainer stands in specified location 

with hands at their sides, beginning 

of training set 

------ Point using two fingers of either 

hand to station desired 

“come” Say come and walk in direction you 

want the animal to go 

“toes” 

“paw” 

“right” 

“left” 

Begin with right arm up parallel to 

body, index and middle fingers 

pointed up. Extend arm straight 

down (palm side down) continuing 

to point both fingers. 

Visual signal for stand; point or 

target foot wanted. 

“up” Cue as for up, trainer body can be 

low 

Animal comes to the front of the 

cage, stands quietly with their eyes 

on the trainer 

Animal moves to the spot and stays 

calmly 

The animal follows and stops 

directly in front of the trainer 

Animal should place both feet 

under the bottom of cage while 

lying down in front of trainer. It 

should be lying still and focused on 

the trainer. 

Cue each foot to right or left, can 

use target or catch less dominant 

foot when opportunity rises; most 

have dominant foot they learn 

easily. 

Same as stand, animal should wait 

while being touched on abdomen 

with pole or wand. 

“touch” Show wand Otter stands on back legs and 

touches target with nose while 

abdomen/kidneys ultrasounded 

through cage mesh. 

“paint” Show painting apparatus Animal grabs paint brush and puts 

paint on canvas. 

“nipple” Target up while standing on hind 

legs. Slowly reach with fingers 

extended, toward otter 

Animal presents chest or abdomen 

against cage mesh for manipulation 

Target placed high on fence Animal climbs fence until all feet off 

the ground and ventrum placed on 

fence 

“water” Use right hand with food in it. Start 

with hand in a fist in front of chest. 

With a sweeping motion, move fist 

up to cage. Arm should be parallel. 

Open hand palm up and out. Tap 

The animal should jump into the 

water to retrieve food.


Water 

^ 

Circle 

^ 

Steady 

^ 

cage with palm to push the food 

into the pool 

“water” Right hand motions towards the 

water 

Animal goes in the water 

“circle” Make a circle with right hand Animal turns in a circle 



“steady” Verbal cue only Used to keep the animal calm 

during tactile body examination 

8.2 Environmental Enrichment 

Environmental enrichment, also called behavioral enrichment, refers to the practice of providing a 

variety of stimuli to the animal’s environment, or changing the environment itself to increase physical 

activity, stimulate cognition, and promote natural behaviors. Stimuli, including natural and artificial objects, 

scents, and sounds, are presented in a safe way for the otters to interact with. Some suggestions include 

providing food in a variety of ways (i.e., frozen in ice or in a manner that requires an animal to solve 

simple puzzles to obtain it), using the presence or scent/sounds of other animals of the same or different 

species, and incorporating an animal training (husbandry or behavioral research) regime in the daily 

schedule. 

It is recommended that an enrichment program be based on 

current information in biology, and should include the following 

elements: goal-setting, planning and approval process, 

implementation, documentation/record-keeping, evaluation, and 

subsequent program refinement. Environmental enrichment 

programs should ensure that all environmental enrichment devices 

(EEDs) are safe and are presented on a variable schedule to 

prevent habituation AZA-accredited institutions must have a formal 

written enrichment program that promotes behavioral opportunities 

(1.6.1). 

Enrichment programs should be integrated with veterinary 

care, nutrition, and otter training programs to maximize the 

effectiveness and quality of animal care provided. AZA-accredited 

institutions must have specific staff members assigned to oversee, 

implement, train, and coordinate interdepartmental enrichment programs (1.6.2). 


(1.6.1) The institution must have a formal 

written enrichment program that promotes 

species-appropriate behavioral 

opportunities. 


(1.6.2) The institution must have a 

specific staff member(s) or committee 

assigned for enrichment program 

oversight, implementation, training, and 

interdepartmental coordination of 

enrichment efforts. 

Development of enrichment ideas should be goal-oriented, proactive, based upon the animal’s natural 

history, individual history and exhibit constraints, and should be integrated into all aspects of their ex-situ 

population management. Providing the appropriate enclosure designs (e.g., land/water ratios, pool/land 

designs), substrates, and furnishings for each otter species are essential components of any enrichment 

program. Enrichment should encourage otters to behave as they would in the wild, as closely as possible. 

Successful enrichment techniques include variation of exhibit schedule or exhibit mates (where 

appropriate only), re-arranging of exhibit furniture/features, complete change of furniture (some of the old 

should always be retained to maintain the animal’s scent and an element of the familiar), scents, sounds, 

toys (natural and artificial), herbs, spices, different substrates for digging/rolling, food items, and novel 

presentation of food items. It is important that enrichment items are not merely thrown in an exhibit and 

allowed to stay for extended periods – an enrichment program is only successful and useful if actively 

managed and constantly reviewed to ensure it encourages natural behaviors. The AAZK Enrichment 

committee provides the following general guidelines about enrichment: 

“The goal of enrichment should be to maximize the benefit while minimizing unacceptable risks. All 

enrichment should be evaluated on three levels: 1) whether the enrichment item itself poses an 

unacceptable risk to the animals; 2) what benefit the animals will derive from the enrichment; and 3) 

whether the manner of enrichment delivery is apt to lead to problems. 

A written plan of action that eliminates the most dangerous risk factors while maintaining the benefits 

of a challenging and complex environment can help animal managers develop a safe and successful 

enrichment program. Keepers should evaluate new and creative enrichment ideas with their managers



and staff from other departments (curatorial, janitorial, maintenance, veterinary, nutritional, etc.) to 

decrease the frequency of abnormal and stereotypic behaviors or low activity levels, and to fine-tune 

enrichment ideas. For enrichment to be safely provided, it is strongly recommended that each institution 

establish enrichment procedures, protocols, and a chain of command that keepers can follow” (AAZK 

Enrichment Committee). 

The AAZK Enrichment Committee also provides an excellent cautionary list for the various types of 

enrichment provided (accessed through www.aazk.org). This list includes key questions that should be 

answered for all enrichment items or programs to assess potential hazards. For example: 

• Can the animals get caught in it or become trapped by it? 

• Can it be used as a weapon? 

• Can an animal be cut or otherwise injured by it? 

• Can it fall on an animal? 

• Can the animal ingest the object or piece of it? Is any part of it toxic, including paint or epoxy? 

• Can it be choked on or cause asphyxiation or strangulation? 

• Can it become lodged in the digestive system and cause gut impaction or linear obstruction? 

• In a multi-species exhibit or other social grouping, could a larger or smaller animal become stuck 

or injured by the object or get hung up on it? 

• Can it destroy an exhibit? 

• If fecal material is used for enrichment, has it been determined to be free from harmful parasites? 

• Is food enrichment included as part of the animals' regular diet in a manner that will reduce the 

risk of obesity? 

• When introducing animals to conspecifics or in a multi-species exhibit, are there sufficient areas 

for them to escape undesirable interactions? 

• Can the manner of enrichment presentation (i.e., one item or items placed in a small area) 

promote aggression or harmful competition? 

• Has browse been determined to be non-toxic? 

• Do animals show signs of allergies to new items (food, browse, substrates)? 

• Does the enrichment cause abnormally high stress levels? 

• Does the enrichment cause stimulation at a high level for extended periods of time that do not 

allow the animal natural down time in the species' normal repertoire (e.g., constant activity for 

public enjoyment when the animal would normally be inactive in its native habitat)?

AAZK Enrichment Committee, Enrichment Caution List 



Dietary Enrichment 

- Food enrichment, if uncontrolled, can lead to obesity, tooth decay and deviation from the normal diet 

can cause nutritional problems. Keepers can consult with the nutritionist or commissary staff to 

determine the best method of introducing novel food items. 

- New food items introduced without analysis may cause colic, rumenitis or metabolic acidosis in 

ungulates. 

- Food items can spoil and cause animal illness if left in the exhibit for extended periods of time. 

Enrichment food items should be removed within a reasonable amount of time to prevent spoilage. 

- Animals can have adverse reactions to toxic plants and chemicals. Keepers should be able to 

correctly discern between toxic and browse plants, ensure that browse is free of fertilizers and 

herbicides and wash plants to remove free ranging bird and animal feces and debris. 

- Foraging or social feedings may give rise to aggression and possible injuries within the animal 

population. 

- Competition for enrichment items may lead to social displacement of subordinate animals. These 

concerns can be minimized by providing enough enrichment to occupy all of the animals within the 

population. 

- Carcass feedings for omnivores and carnivores may be hazardous if the source of the carcass is not 

determined and appropriate precautions taken. Diseased animals, chemically euthanized animals or 

those with an unknown cause of death are not appropriate for an enrichment program. Freezing the 

carcasses of animals that are determined to be safe to feed to exhibit animals can help minimize the 

risk of parasitism and disease. Providing enough carcasses in group feedings can minimize 

competition and aggression within an exhibit. 

- Carefully introducing a group of animals to the idea of social feedings can be done by moving carcass 

pieces closer together at each feeding until the animals are sharing one carcass. This can allow 

social carnivores to exhibit normal dominance posturing while minimizing the possibility of 

aggression. During live feedings, prey animals may fight back. Care should be taken to ensure such 

prey can only inflict superficial wounds on zoo animals. 

- Cage furniture may interrupt flight paths or entangle horns and hooves if poorly placed. Careful 

planning can prevent this. 

- If unsecured, some items may fall on an animal or be used as a weapon and cause injuries. 

- If position is not thoughtfully considered, limbs and apparatus may provide avenues for escape or 

may block access into exhibit safety zones, leaving subordinate animals feeling trapped and 

vulnerable. 

- Animals that crib or chew wood should be provided with non-toxic limbs and untreated wood furniture. 

- Water features should be tailored to the inhabitants to prevent drowning and ensure that animals 

such as box turtles can right themselves if they flip over on their backs. 

- Animals can be injured in filtration systems if water intake areas are not protected. 

- Substrates should provide adequate traction and not cause an intestinal impaction if ingested. 

- Caution should be exercised when ropes, cables, or chains are used to hang or secure articles to 

prevent animals from becoming entangled. Generally, the shortest length possible is recommended. 

Chain can be covered with a sheath such as PVC pipe; swivels can be used to connect the chain to 

the enrichment item to minimize kinking. 

Olfactory Enrichment 

- Scents from different animals or species can lead to aggression if there is an assertion of dominant 

animals or subordinate animals attempting to use enrichment to advance their status in the hierarchy. 

- Animal feces used for olfactory enrichment should be determined to be parasite free through fecal 

testing and as with other animal by-products such as feathers, sheds, wool and hair, come from only 

healthy animals. Many of these items can be autoclaved for sterilization. 

- Perfumes can be overwhelming to some animals (and keepers) and are therefore best used in open, 

ventilated areas. 

- Some spices may be too strong or toxic to some animals.



Auditory Enrichment 

- When provided with audio enrichment, animals may be less threatened by deflected sounds rather 

than those directed at the animals. 

- Some animals may have adverse reactions to recordings of predator calls and should be closely 

observed when this type of enrichment is provided. 

- Providing the animals with an option for escape or the means to mobilize for confrontation when 

predator calls are played can lessen the stress of this type of enrichment and allow the animals to 

investigate the sounds and their environment over a period of time. 

Manipulable Enrichment 

- Individual parts or enrichment devices may be swallowed resulting in choking or asphyxiation. 

- If ingested, indigestible enrichment items may cause a gut impaction or linear obstruction. 

- Broken items may have sharp edges that can cut an animal. Only items that are appropriate for the 

species should be provided. For example, some devices will hold up to the play of a fox but not a wolf 

- When building or designing enrichment items from wood, it may be wise to use dovetail cuts and glue 

rather than screws and nails. Rounded corners and sanded edges can prevent the animals from 

getting splinters. 

- Many paints and other chemicals are toxic if eaten. When providing enrichment involving paint or 

other chemicals, only non-toxic items should be used. 

- If used, destructible items such as cardboard boxes and paper bags should be free of staples, tape, 

wax, strings or plastic liners. In general the Otter SSP advises against using these items. 

Factors that should be considered when determining how often behavioral or environmental 

enrichment is offered include the species and individual(s) involved, as well as the physical 

characteristics of the exhibit. Large, complex exhibits with appropriate enclosure designs, substrates, 

and furnishings may offer ample opportunities for animals to exercise natural behaviors with infrequent 

enrichment (once daily). Other exhibits or individuals may require more frequent enrichment (multiple 

times per day). Husbandry staff should monitor all individuals in an exhibit and structure an enrichment 

schedule for the needs of those animals, providing them with opportunities several times a day to interact 

positively with their environment. Enrichment should never be offered on a regular schedule, instead 

times, items, and delivery methods should be rotated so there is always an element of novelty associated 

with each item or activity. It is important to note that the provision of well-designed, complex environments 

is the foundation of a successful enrichment program. This is particularly true for some of the more 

sensitive otter species such as P. brasiliensis, but applies to all of the otter species due to their inquisitive 

nature and high-activity level. 

More Information: Appendix K provides a list of enrichment initiatives used at several institutions 

housing mustelids/otters. All enrichment items should be approved by the appropriate management staff, 

including the veterinarian, curator, horticulturist, and/or nutritionist. Appendix L provides a list of resources 

for enrichment and training. Institutions working with P. brasiliensis should consult the International Giant 

Otter Studbook Husbandry and Management Information and Guidelines (Sykes-Gatz 2005) for further 

information on the importance of exhibit design for this species and additional enrichment information. 

8.3 Staff and Animal Interactions 

Animal training and environmental enrichment protocols and techniques should be based on 

interactions that promote safety for all involved. Otters are easily trained and work well for positive 

reinforcement. Trained management and veterinary care behaviors include: weighing, crating, foot 

inspection, tooth inspection, injections, abdominal presentation and palpation, and tail inspections. It 

should be kept in mind that otters are capable of inflicting severe bites, particularly sexually mature 

females, and have been known to turn on their trainers. In general, otters should be trained in a protected 

contact situation (i.e., keeper and animal should be separated by a mesh barrier). There are some 

species (A. cinereus) or cases (L. canadensis particularly males) where an institution may feel that 

protected contact training may not be called for, but these decisions should be carefully evaluated on an 

ongoing basis. 

Keeper safety should be kept in mind when designing otter exhibits. Animals should be shifted off 

exhibit for cleaning, maintenance, etc. This is particularly important for P. brasiliensis, as they can be



dangerous. Animal safety should be considered and exhibits constructed, as far as possible, that prohibit 

the public from throwing potentially harmful items or food into the animals’ space. 

8.4 Staff Skills and Training 

Staff members should be trained in all areas of animal behavior management. Funding should be 

provided for AZA continuing education courses, related meetings, conference participation, and other 

professional opportunities. A reference library appropriate to the size and complexity of the institution 

should be available to all staff and volunteers to provide them with accurate information on the behavioral 

needs of the otters with which they work. 

The following skills are recommended for all animal caretakers involved in the management of otters 

of ex-situ populations: 

• Keepers and managers should have an in-depth understanding of the species’ natural history and 

the individual’s history. 

• Keepers and managers should have an in-depth understanding of the individual’s behaviors, an 

understanding of the function of those behaviors, and the ability to describe those behaviors 

orally and in writing. 

• Keepers should be able to recognize signs of illness and injury in the otter species they are 

working with and to communicate those signs orally or in writing to managers and veterinarians. 

• Keepers should be able to accurately assess the appropriate level of cleanliness and safety of the 

animal’s exhibit, holding area, and food-prep area. 

• Keepers should have the skills to safely capture or restrain the otter species in question. 

• Keepers should have some understanding of the species’ natural diet and foraging style. 

• Keepers and managers should have an understanding of enrichment concepts and have a 

commitment to consistently enhance the environments of the species in their care. 

• Keepers should understand the concepts of animal learning and training, be able to use a variety 

of techniques (e.g., habituation, operant conditioning) to train the animals under their care, and to 

create a training plan (identifying training steps, cues, and criteria). See www.animaltraining.org 

for additional information. 

• Managers should understand the concepts of animal learning and training, be able to coach 

keepers in all aspects of training, review their training plans, look for consistency among keepers 

in their training techniques, and help their teams prioritize training, enrichment, and other 

husbandry goals. 

• Keepers and managers should have an understanding of the enclosure conditions and husbandry 

practices needed to maintain the otters’ physical and behavioral health, as well as to promote a 

successful pup-rearing environment.



Chapter 9. Program Animals 

9.1 Program Animal Policy 

AZA Policies: AZA recognizes many public education and, ultimately, conservation benefits from 

program animal presentations. AZA’s Conservation Education Committee’s Program Animal Position 

Statement (Appendix D) summarizes the value of program animal presentations. 

For the purpose of this policy, a program animal is described as an animal presented either within or 

outside of its normal exhibit or holding area that is intended to have regular proximity to or physical 

contact with trainers, handlers, the public, or will be part of an ongoing conservation education/outreach 

program. 

Program animal presentations bring a host of responsibilities, including the welfare of the animals 

involved, the safety of the animal handler and public, and accountability for the take-home, educational 

messages received by the audience. Therefore, AZA requires all accredited institutions that give program 

animal presentations to develop an institutional program animal policy that clearly identifies and justifies 

those species and individuals approved as program animals and details their long-term management plan 

and educational program objectives. 

AZA’s accreditation standards require that the conditions and 

treatment of animals in education programs must meet standards 

set for the remainder of the animal collection, including speciesappropriate 

shelter, exercise, sound and environmental 

enrichment, access to veterinary care, nutrition, and other related 

standards (1.5.4). In addition, providing program otters with 

options to choose among a variety of conditions within their 

environment is essential to ensuring effective care, welfare, and 

management. Some of these requirements can be met outside of 

the primary exhibit enclosure while the otter is involved in a 

program or is being transported. For example, housing may be 

reduced in size compared to a primary enclosure as long as the 

otter’s physical and psychological needs are being met during the 

program; upon return to the facility the otter should be returned to 

its housing as described above. 


(1.5.4) A written policy on the use of live 

animals in programs should be on file. 

Animals in education programs must be 

maintained and cared for by trained staff, 

and housing conditions must meet 

standards set for the remainder of the 

animal collection, including speciesappropriate 

shelter, exercise, social and 

environmental enrichment, access to 

veterinary care, nutrition, etc. Since some 

of these requirements can be met outside 

of the primary enclosure, for example, 

enclosures may be reduced in size 

provided that the animal’s physical and 

psychological needs are being met. 

Otter SSP Program Animal Recommendations: A. cinereus and occasionally L. canadensis are used 

in training shows however, otters are not recommended as animal handling or off-site education animals. 

Conservation Messages: Otters are excellent conservation and wildlife ambassadors; they are 

appealing to the public, active, and represent well the issues faced by many of the small carnivores. 

African spotted-necked otter: “Spotted-Necked Otters are very aquatic and require permanent water 

sources with high fish densities. They prefer larger rivers, lakes, and swamps with open areas of water. 

They appear to only make use of fresh water habitats. Because they mainly hunt by sight, they need 

clear, unpolluted water where there are numerous small fish or fish, crabs and frogs. Long reeds, grass, 

and bushes are essential to provide cover; holes or other shelters are also needed. The most suitable 

habitat is the large fish-rich African lakes and the deep, clear areas of the Botswana Okavango. 

“The distribution is large, but with some local declines. It occurs in all countries south of Sahara, from 

Senegal to Ethiopia and south to the Cape provinces where there is suitable habitat. 

“The main threats throughout the range are habitat destruction by land drainage or pollution in 

response to increasing human population density and direct persecution as competitors for fish. There is 

some hunting for bushmeat and ceremonial practices. In some lakes, introduced large fish such as Nile 

Perch out-compete the small fish, which comprise the otters' historic food base, reducing prey availability. 

Although international and national level legal protection is in place, enforcement is needed. There is 

a need for increasing local awareness of the species (IUCN Otter Specialist Group).” 

The population is considered to be decreasing, and there are some indications that pressure from 

traditional medical uses, bushmeat consumption, and persecution as competitors for fish on some 

populations (Lake Victoria) may be increasing (J.Reed-Smith, personal knowledge).



Asian small-clawed otter: “Small-clawed otters prefer shallow water with a good food supply and 

moderate to low bank side vegetation. They demonstrate a high climatic and trophic adaptability, 

occurring from tropical coastal wetlands up to mountain streams. They make use of freshwater and peat 

swamp forests, rice fields, lakes, streams, reservoirs, canals, drainage ditches, rice paddies, mangroves, 

tidal pools, and along the coastline. In mountainous areas, they frequent swift-flowing forest streams with 

rocks and boulders. Their preferred food is crustaceans and mollusks. Across much of their range they 

are sympatric with Eurasian Otters (Lutra lutra), Smooth-Coated Otters (Lutrogale perspicillata) and 

Hairy-Nosed Otters (Lutra sumatrana), and there is clear evidence of niche separation between the 

species. 

Although the species' range appears large, in the last decade actual distribution has shrunk, 

especially in the west, compared to historical records. They are currently found from the Himalayan 

foothills of Himachal Pradesh eastward throughout south Asia, extending up to Philippines and down 

through Indonesia. A small isolated subpopulation has been reported from southern Indian hill ranges of 

Coorg (Karnataka), Ashambu, Nilgiri and Palni hills (Tamil Nadu), and some places in Kerala. They were 

formerly found in Sri Lanka, but their current status there is unknown. The only areas in which these 

animals are known to be common today are Peninsular Malaysia, especially in Kedah, and in the western 

forests and southern marshes of Thailand. 

The main threats throughout Asia are habitat destruction because of deforestation (loss of the smaller 

hill streams), agriculture (especially tea and coffee plantations in India, draining of peat swamp forests, 

and destruction of coastal mangroves for aquaculture), and settlement. Water courses are being polluted 

with pesticides from plantations and other intensive agriculture and heavy metals, affecting the gillfeeders 

on which this species depends. This interferes directly with otter physiology. Prey biomass is also 

being reduced by overexploitation, and the vast aquaculture industry regards otters as pests and 

persecutes them directly. 

Although international and national level legal protection is in place, local legislation is needed. The 

impact of protection measures on livelihoods needs to be assessed and answered. Habitat protection and 

interpopulation corridors need to be established. Research on all aspects of this species biology and 

ecology is needed (IUCN Otter Specialist Group).” The population is considered to be decreasing. 

Giant otter: “The wild population is estimated to have a total population of 1,000 to 5,000 individuals. In 

the past, giant otters were frequently hunted for their fur. This trade in giant otter pelts is one of the 

primary reasons giant otters are endangered in the wild. Because this species is active during the day, 

very vocal, and not afraid to approach humans, they were easy to hunt. Much of the population became 

decimated until efforts were finally made to protect them in the 1970’s. 

Although they still face other serious threats, habitat destruction and degradation, poaching, and 

unmanaged tourism are the primary threats faced by giant otter today. The areas in South America where 

the giant otter lives are rapidly being destroyed and degraded by logging, mining, exploitation of fossil 

fuels and hydroelectric power (dams), river and land pollution, and over-fishing. Some giant otter cubs are 

still being taken from the wild illegally to be kept as pets and they usually die in the hands of 

inexperienced caretakers. Tourists can disturb giant otters when they are rearing cubs. This can have a 

negative effect on how successfully parents rear their litters (IUCN Otter Specialist Group).” The 

population is considered to be decreasing. 

North American river otter: This species is also referred to as the Nearctic otter. However, whatever you 

call it, the river otter represents a North American conservation success story. From a historic high when 

their range extended throughout most of North America, river otter populations fell until: 

“During the late 1800’s and early 1900’s, the synergistic effect of wetland destruction, 

pollution, and overexploitation for furs was devastating to North American river otter 

populations. Additional otter losses were due to road kills, accidental drowning in fishing nets 

and ‛incidental take during beaver trapping’.” (Foster-Turley et al. 1990) 

By the 1970’s, Nilsson & Vaughn (1978) estimated that the river otter was found in only 33% of its 

former range. They listed the causes of this as intensive trapping, pollution, destruction of habitat by 

clearing land, draining marshes, and channelizing streams. However, since 1976 over 4,000 otters have 

been reintroduced in 21 states and provinces throughout their former North American range (including 

Canada).



As a result of conservation and reintroduction measures the river otter has reoccupied much of its 

former range; this species it is still considered locally vulnerable, endangered, or extinct in some states 

and provinces. Many states and provinces where the populations were never threatened or have 

recovered allow a sustainable harvest of this species for fur. While “current harvest strategies do not pose 

a threat to maintaining otter populations, harvest may limit expansion of otter populations in some areas. 

Oil spills present a localized threat to otter populations, especially in coastal areas. Water pollution and 

other degradation of aquatic and wetland habitats may limit distribution of otters and pose long-term 

threats if enforcement of water quality standards are not maintained and enforced. Acid drainage from 

coal mines is a persistent water quality problem in some areas that eliminates otter prey and thereby 

inhibits recolonization or expansion of otter populations. Recently, there has been discussion of the longterm 

genetic consequences of reintroduction projects on remnant otter populations (Serfass et al. 1998). 

The threat of disease to wild otter populations is poorly understood and has received little study (Serfass 

et al. 1995). Similarly, many perceived threats to otters such as pollution and habitat alterations have not 

been rigorously evaluated. Additional research is needed to clearly delineate the impact that various 

forms of water pollution, agricultural and other development along riparian habitats, industrial and housing 

development in coastal areas, cumulative impacts related to loss or alterations of wetlands, large flood 

control structures, and interactions that these and other factors have on otter populations. Threats to otter 

populations in North America vary among regions and are influenced by type, distribution, and density of 

aquatic habitats and characteristics of human activities (IUCN Otter Specialist Group).” 

Recently, concern has been growing that river otters are being demonized as voracious eaters of 

sport-fish species; this position has been used to justify elimination of river otters from some watersheds 

to placate special interest groups. Some research has been conducted on the percentage of sport fish 

species taken by river otter that indicates the true impact is not as great as claimed by many (Hamilton, 

1999 & unpublished 2004); See Appendix L. The impact of river otter on sport-fish populations and the 

growing characterization of the river otter as the primary cause of localized falling fish populations is 

something that should be researched and monitored. 

9.2 Institutional Program Animal Plans 

AZA’s policy on the presentation of animals is as follows: AZA is dedicated to excellence in animal 

care and welfare, conservation, education, research, and the presentation of animals in ways that inspire 

respect for wildlife and nature. AZA’s position is that animals should always be presented in adherence to 

the following core principles: 

• Animal and human health, safety, and welfare are never compromised. 

• Education and a meaningful conservation message are integral components of the presentation. 

• The individual animals involved are consistently maintained in a manner that meets their social, 

physical, behavioral, and nutritional needs. 

AZA-accredited institutions which have designated program animals are required to develop their own 

Institutional Program Animal Policy that articulates and evaluates the program benefits (see Appendix E 

for recommendations). Program animals should be consistently 

maintained in a manner that meets their social, physical, 

behavioral, and nutritional needs. Education and conservation 

messaging must be an integral component of any program 

animal demonstration (1.5.3). 

Animal care and education staff should be trained in otterspecific 

handling protocols, conservation and education 

messaging techniques, and public interaction procedures. These 

staff members should be competent in recognizing stress or 

discomfort behaviors exhibited by the program otters and should 

be able to address any safety issues that arise. 

Program animals that are taken off zoo or aquarium grounds 

for any purpose have the potential to be exposed to infectious 

agents that could spread to the rest of the institution’s healthy 

population. AZA-accredited institutions must have adequate 

protocols in place to avoid this (1.5.5). 


(1.5.3) If animal demonstrations are a part 

of the institution’s programs, an education 

and conservation message must be an 

integral component. 


(1.5.5) For animals used in offsite 

programs and for educational purposes, 

the institution must have adequate 

protocols in place to protect the rest of the 

collection from exposure to infectious 

agents.

Careful consideration must be given to the design and size of 

all program animal enclosures, including exhibit, off-exhibit 

holding, hospital, quarantine, and isolation areas, such that the 

physical, social, behavioral, and psychological needs of the 

species are met and species-appropriate behaviors are facilitated 

(10.3.3; 1.5.2). 

Animal transportation must be conducted in a manner that is 

lawful, safe, well-planned and coordinated, and minimizes risk to 

the animal(s), employees, and general public (1.5.11). 

9.3 Program Evaluation 

AZA-accredited institutions which have Institutional Program 

Animal Plan are required to evaluate the efficacy of the plan 

routinely (see Appendix E for recommendations). Education and 

conservation messaging content retention, animal health and 

well-being, guest responses, policy effectiveness, and 

accountability and ramifications of policy violations should be 

assessed and revised as needed. 




(10.3.3) All animal enclosures (exhibits, 

holding areas, hospital, and 

quarantine/isolation) must be of a size 

and complexity sufficient to provide for 

the animal’s physical, social, and 

psychological well-being; and exhibit 

enclosures must include provisions for the 

behavioral enrichment of the animals. 


(1.5.2) Animals should be displayed, 

whenever possible, in exhibits replicating 

their wild habitat and in numbers sufficient 

to meet their social and behavioral needs. 

Display of single specimens should be 

avoided unless biologically correct for the 

species involved. 


(1.5.11) Animal transportation must be 

conducted in a manner that is safe, wellplanned 

and coordinated, and minimizes 

risk to the animal(s), employees, and 

general public. All applicable local, state, 

and federal laws must be adhered to.

10.1 Known Methodologies 

AZA believes that contemporary animal management, 

husbandry, veterinary care and conservation practices should be 

based in science, and that a commitment to scientific research, 

both basic and applied, is a trademark of the modern zoological 

park and aquarium. AZA-accredited institutions have the 

invaluable opportunity, and are expected, to conduct or facilitate 

research both in in-situ and ex-situ settings to advance scientific 

knowledge of the animals in our care and enhance the 

conservation of wild populations. For otters, this knowledge might 

be achieved by AZA Small Carnivore Taxon Advisory Group 



Chapter 10. Research 

(TAG) or AZA Otter Species Survival Plan® (SSP) Program sponsored research, conducting original 

research projects, affiliating with local universities, and/or employing staff with scientific credentials (5.3). 

Research investigations, whether observational, behavioral, physiological, or genetically based, 

should have a clear scientific purpose with the reasonable expectation that they will increase our 

understanding of the otter species being investigated and may provide results which benefit the health or 

welfare of otters in wild populations. Many AZA-accredited institutions incorporate superior positive 

reinforcement training programs into their routine schedules to 

facilitate sensory, cognitive, and physiological research 

investigations and these types of programs are strongly 

encouraged by the AZA. 

AZA-accredited institutions are required to have a clearly 

written research policy that identifies the types of research being 

conducted, methods used, staff involved, evaluations of the 

projects, the animals included, and guidelines for the reporting or 

publication of any findings (5.2). Institutions must designate e a 

qualified individual designated to oversee and direct its research 

program (5.1). If institutions are not able to conduct in-house 

research investigations, they are strongly encouraged to provide 

financial, personnel, logistical, and other support for priority 

research and conservation initiatives identified by Taxon Advisory 

Groups or Species Survival Plans®. 


(5.3) Institutions should maximize the 

generation of scientific knowledge gained 

from the animal collection. This might be 

achieved by participating in AZA 

TAG/SSP sponsored research when 

applicable, conducting original research 

projects, affiliating with local universities, 

and/or employing staff with scientific 

credentials. 


(5.2) Institutions must have a written 

policy that outlines the type of research 

that it conducts, methods, staff 

involvement, evaluations, animals to be 

involved, and guidelines for publication of 

findings. 


(5.1) Research activities must be under 

the direction of a person qualified to make 

informed decisions regarding research. 

The AZA Otter SSP, which falls under the Small Carnivore TAG, is the AZA entity tasked with 

recommending species for management by member institutions. The IUCN/SSC Otter Specialist Group 

(OSG) serves as an international focus of information sharing for these species. Information on current 

and past field work can be found at the OSG web site, www.otterspecialistgroup.org. Several universities 

(e.g. Frostburg State University, University of Wyoming) have professors specializing in otter research. 

However, situations change and students interested in pursuing work with otters always should research 

current specialists working in the area and institutions with which they are affiliated. The AZA Otter SSP is 

compiling a list of institutions involved in or supporting otter research and conservation work. This 

information will become available in future versions of this document. For input on ongoing research or 

areas requiring further investigation please refer to the OSG web site or contact the AZA Otter SSP Chair. 

Effective Research Methodologies: All sound research approaches should be viable for use on otters, 

as long as they are not too invasive, require extensive surgery, or cause pain or discomfort. The following 

methodologies are routinely used by researchers looking at otters: 

• Behavioral observation 

• Latrine surveys 

• Fecal hormone analysis represents an invaluable tool for assessing the reproductive status of 

individuals and populations in a completely noninvasive manner. 

• Fecal DNA analysis



10.2 Future Research Needs 

This Otter Care Manual is a dynamic document that will need to be updated as new information is 

acquired. Knowledge gaps have been identified throughout the Manual and a general list is included in 

this section to promote future research investigations. Knowledge gained from areas will maximize AZAaccredited 

institutions’ capacity for excellence in otter care and welfare as well as enhance conservation 

initiatives for all species. 

Specific Areas of Ex-situ Population Research Needed by Manual Heading: 

Chapter 1: Ambient Environment 

1.1 Water temperature: More detailed research is required into optimal water temperature levels for the 

tropical otter species. 

1.2 Light: The AZA Small Carnivore TAG is unaware of any hard data on the impact of light intensity on 

otter health or reproduction; this should be investigated in the future. Similarly, there are no available data 

on possible deleterious effects of less than full spectrum light on a long-term basis. Health data collected 

by institutions housing otters in environments with different light intensities and spectrums would be a 

useful foundation for this research. 

1.3 Air: Pupping dens may well need higher rates of air exchange in order to maintain air quality and/or 

low humidity. It should be noted, however, that no work has been done specifically targeting air change 

rates for otter exhibits or dens. 

1.3 Water - Coliform: There are no standards for coliform yet established for fresh-water otter pools. At 

this time, it is suggested that coliform levels be maintained at or lower than levels established for rescued 

pinnipeds by NOAA. 

1.3 Water - pH Levels: Further research is needed into the impact, if any, of pH on otters. While research 

is desirable it is not recommended on these wildlife species and instead caution should be exercised 

when using chlorine in otter pools. 

1.4 Sound: While there is no evidence that low level background noise is disruptive to otters, loud noises 

can be frightening to them, and high-pitched, long-term noise should be avoided. Otters’ hearing is 

considered to be good but nothing is known definitively about their hearing acuity or frequency ranges 

heard. Both of these are areas needing further research. 

Chapter 3: Transport 

3.2 Transport temperature: Identifying appropriate transport temperatures for L. maculicollis requires 

further research. Current recommendations are based on A. cinereus, but it is not known whether this is 

suitable. 

Chapter 4: Social Environment 

4.1 Single-sexed groups: Information on the success of single-sexed groups for A. capensis is unknown 

at this time; further research is required. Information on the maintenance of family groups of L. 

maculicollis over the long term has not been well documented; this should be monitored to assist in future 

recommendations for this species. Information from institutions attempting this type of social grouping 

should be shared with the AZA Small Carnivore TAG. 

Chapter 5: Nutrition 

5.1 Seasonal changes in nutritional needs: Some institutions report seasonal changes in appetite of 

some otters, but not in the majority of animals. Further research in this area, and in changing seasonal 

nutritional requirements for otters, is required. 

5.1 Nutritional related diseases: Further research on nutritional requirements of otters and nutrition 

related diseases. 

5.2 Sample diets: The one best diet for any of the otters of ex-situ populations has not been found and 

requires further research. Identifying health issues associated with the provision of different diets should 

continue, and data shared with the AZA Small Carnivore TAG.



5.3 Nutrition evaluation: The AZA Otter SSP is currently beginning work on a body-condition matrix that 

can be used to help assess proper weight and condition for otters. At this time there are no known tools 

for performing clinical nutritional evaluations of otters; this would be a useful area for future research. 

Chapter 6: Veterinary Care 

6.7 Management of diseases and disorders: Little information on common diseases and disorders for 

A. capensis and L. maculicollis is available, and more research is required for these species. Institutions 

housing these species should record all health issues seen in these species so that a database can be 

created on health issues and concerns. 

Chapter 7: Reproduction 

7.1 Reproductive physiology and behavior: Some facilities have reported a small amount of estrusassociated 

bleeding from the vulva in L. canadensis, while others have not seen this; additionally, 

previous studies attempting to identify behavioral changes associated with estrus were unsuccessful. 

Information on female estrus behavior would be a helpful area of research. These are areas that require 

further research and can be achieved through simple observational research during estrus for animals in 

this species. 

7.1 Reproductive physiology and behavior: In both NARO and ASCO, additional research is needed to 

improve endocrine monitoring of estrogen metabolites to further address these questions about ovarian 

cyclicity and ovulatory mechanisms. 

7.1 Reproductive physiology and behavior: Research utilizing techniques to identify reproductive state 

in these species is ongoing. At this time, it appears that ELISA protocols for testing hormonal secretions 

in fecal samples are successful in determining pregnancy in Asian small-clawed and North American river 

otters (H.Bateman, unpublished data). The reproductive physiology advisor for the AZA Otter SSP, 

should be contacted for more information. 

7.1 Reproductive physiology and behavior: Further study is required to clarify if there is a genetic 

component to the seasonal regulation of estrus in females and testosterone production in NARO males. 

7.2 Pregnancy and parturition: Pseudopregnancy has been reported for most otter species and is an 

area that requires further research 

7.3 Pup development: At this time there is no information available on pup development of A. capensis; 

additional data on the development of L. maculicollis pups is also required. Institutions housing these 

species (A. capensis is a phase-out but still maintained by a few member institutions) should set-up an 

observational research study for this species when reproduction is attempted. 

7.4 Nursery groups: Nursery groups are not reported for L. maculicollis in the wild, but further field 

research would help to determine if this aspect of parental care is applicable or appropriate for ex-situ 

population management of this species. 

7.5 Hand-rearing: More research is required to determine body temperature norms for young of all otter 

species. This information should be collected by all facilities hand-rearing otter pups and submitted to the 

AZA Otter SSP and Small Carnivore TAG. 

7.5 Contraception: Research on the effects that GnRH agonists have on future reproductive abilities 

when provided to prepubertal otters is also needed, as current research is based on studies using 

domestic cats. 

Chapter 8: Behavioral Management 

8.1 Training: Training animals to station may be beneficial when attempting introductions, but this has 

not been tried with any of the otters. 

Other Areas of Research: 

Giant otters: The following priorities were established for the giant otter during the 2004 meeting of the 

IUCN/SSC Otter Specialist Group:



1. To continue the assessment of predator-prey relationships, including conflicts with subsistence 

and commercial fishermen. 

2. To evaluate the positive and negative impact of tourism on different habitats and implement 

management guidelines in order to maximize the benefits. 

3. To encourage the development of a long-term research and conservation project in the Llanos of 

Venezuela or Colombia. 

4. To undertake collaborations between field scientists, zoos, and genetic labs to evaluate the 

potential use of genetic analysis tools in giant otter research. 

Field Research: Additional research is needed to clearly delineate the impact that various forms of water 

pollution, agricultural and other development along riparian habitats, industrial and housing development 

in coastal areas, cumulative impacts related to loss or alterations of wetlands, large flood control 

structures, and interactions that these and other factors have on otter populations (N.A. river otter) 

The impact of river otter on sport-fish populations and the growing characterization of the river otter as the 

primary cause of localized falling fish populations is something that should be researched and monitored. 

General Areas of Research Needed: 

• Nutritional needs to include nutrition related diseases 

• Reproductive research – reproductive physiology research is ongoing (Helen Bateman, 

C.R.E.W.) and should continue for all otter species. 

• Veterinary issues – to include efficacy of vaccines, titers, and common causes of death in ex-situ 

populations. All otters should be necropsied and results sent to the veterinary advisor. 

• Status in the wild – institutions should assist the AZA Otter SSP in raising awareness and funds 

to adequately assess the status of wild otter populations of all species. 

• Ex-situ population behavior studies – to assess extent and type of stereotypes observed in the 

ex-situ population, to assess breeding/parturition behavior; to assess optimum group size and 

composition for otters of ex-situ populations.



Acknowledgements 

The AZA Otter SSP and animal care guideline coordinator for the SCTAG would like to express 

thanks to the many AZA members and non-AZA members who contributed their expertise to the creation 

of these guidelines. We also acknowledge the commitment to quality care of zoo and aquarium animals 

demonstrated by the extraordinary support received from Columbus Zoo and Aquarium, Maryland Zoo in 

Baltimore, and Cincinnati Zoo & Botanical Garden/C.R.E.W. All three institutions supported the creation 

of this document by contributing extensive amounts of valuable personnel time and expertise. Dr. Joseph 

Barber assisted immeasurably throughout the early creation and first edition of this document; his help 

and patience are greatly appreciated. 

It is important to note that the recommendations contained in this manual are fluid and may need to 

be modified to offer best practice care to particular individual animals. The most important thing to 

remember is to know your animals and pay attention to their individual needs. Questions regarding any of 

the information contained in this manual may be directed to Jan Reed-Smith (lontracat@live.com) or 

Dusty Lombardi (dusty.lombardi@columbuszoo.org). 

For additional information on otters, otter conservation, otter research, and otter care we recommend 

the IUCN/SSC Otter Specialist Group web site at: www.otterspecialistgroup.org or the International Otter 

Survival Fund at: www.otter.org.



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Corredor, G.: Curator, Cali Zoo. Cali, Columbia. 

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Grant, Kerrin: Dept. of Forest, Range and Wildlife Sciences Utah State University Logan, Utah, 

zoonutrition@msn.com. 

Harshaw, Chip: AZA Otter SSP member.



Kollias, George: Cornell University School of Veterinary Medicine. 

Landis, Robert: Wyoming, wildlife filmmaker. 

Lengel, Kim: Philadelphia Zoo, Vice President of Conservation. 

Lent, Cheryl: Indianapolis Zoo, Asian small-clawed otter keeper, CLent@Indyzoo.com. 

Lombardi, Dusty: Columbus Zoo and Aquarium, AZA Otter SSP Chair, dusty.lombardi@columbuszoo.org. 

Maher, Sue: Disney’s Animal Kingdom, SCTAG Representative, sue.maher@disney.com. 

Myers, Gwen: Columbus Zoo and Aquarium. Otter SSP Vet. Advisor, Gwen.meyers@columbuszoo.org. 

Meyerson, Randi: Toledo Zoological Garden. AZA Otter SSP., randi@toledozoo.org. 

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Specialist Group. jan.smith@columbuszoo.org. 

Shannon, Scott: Field researcher, North American otter, jss@riverotters.net. 

Spelman, Lucy: Researcher former Director National Zoo, no current contact 

Stark, Beth: Curator of Behavioral Husbandry and Research, Toledo Zoo. Beth.stark@Toledozoo.org. 

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Otter Specialist Group (sheilasykes@hotmail.com) 

Varsik, Alan: Santa Barbara Zoo, General Curator, avarsik@sbzoo.org. 

Walz, Deanna: Utah’s Hogle Zoo, Behavioral Enrichment Coordinator/Sr. Keeper. 

2enrichntrain@hoglezoo.org. 

Willison, R.: Spotted-necked otter keeper, San Diego Zoo, zooroach1@earthlink.net. 

Wilson, Claudia: Wildlife Conservation Society, Bronx, Collection Manager, Mammal Dept. AZA Otter 

SSP, zooroach1@earthlink.net. 

Yoxon, Grace: International Otter Survival Fund. Scotland. Iosf2@aol.com.



Appendix A: Accreditation Standards by Chapter 

The following specific standards of care relevant to Lutrinae are taken from the AZA Accreditation 

Standards and Related Policies (AZA 2009) and are referenced fully within the chapters of this animal 

care manual: 

Chapter 1 

(1.5.7) The animal collection must be protected from weather detrimental to their health. 

(10.2.1) Critical life-support systems for the animal collection, including but not limited to plumbing, 

heating, cooling, aeration, and filtration, must be equipped with a warning mechanism, and 

emergency backup systems must be available. All mechanical equipment should be under a 

preventative maintenance program as evidenced through a record-keeping system. Special 

equipment should be maintained under a maintenance agreement, or a training record should show 

that staff members are trained for specified maintenance of special equipment. 

(1.5.9) The institution must have a regular program of monitoring water quality for collections of fish, 

pinnipeds, cetaceans, and other aquatic animals. A written record must be maintained to document 

long-term water quality results and chemical additions. 

Chapter 2 

(1.5.2) Animals should be displayed, whenever possible, in exhibits replicating their wild habitat and in 

numbers sufficient to meet their social and behavioral needs. Display of single specimens should be 

avoided unless biologically correct for the species involved. 

((10.3.3) All animal enclosures (exhibits, holding areas, hospital, and quarantine/isolation) must be of a 

size and complexity sufficient to provide for the animal’s physical, social, and psychological wellbeing; 

and exhibit enclosures must include provisions for the behavioral enrichment of the animals. 

(11.3.1) All animal exhibits and holding areas must be secured to prevent unintentional animal egress. 

(11.3.6) Guardrails/barriers must be constructed in all areas where the visiting public could have contact 

with other than handleable animals. 

(11.2.3) All emergency procedures must be written and provided to staff and, where appropriate, to 

volunteers. Appropriate emergency procedures must be readily available for reference in the event of 

an actual emergency. These procedures should deal with four basic types of emergencies: fire, 

weather/environment; injury to staff or a visitor; animal escape. 

(11.6.2) Security personnel, whether staff of the institution, or a provided and/or contracted service, must 

be trained to handle all emergencies in full accordance with the policies and procedures of the 

institution. In some cases, it is recognized that Security personnel may be in charge of the respective 

emergency (i.e., shooting teams). 

(11.2.4) The institution must have a communication system that can be quickly accessed in case of an 

emergency. 

(11.2.5) A written protocol should be developed involving local police or other emergency agencies and 

include response times to emergencies. 

(11.5.3) Institutions maintaining potentially dangerous animals (sharks, whales, tigers, bears, etc.) must 

have appropriate safety procedures in place to prevent attacks and injuries by these animals. 

Appropriate response procedures must also be in place to deal with an attack resulting in an injury. 

These procedures must be practiced routinely per the emergency drill requirements contained in 

these standards. Whenever injuries result from these incidents, a written account outlining the cause 

of the incident, how the injury was handled, and a description of any resulting changes to either the 

safety procedures or the physical facility must be prepared and maintained for five years from the 

date of the incident. 

Chapter 3 

(1.5.11) Animal transportation must be conducted in a manner that is safe, well-planned and coordinated, 

and minimizes risk to the animal(s), employees, and general public. All applicable local, state, and 

federal laws must be adhered to. 

Chapter 5 

(2.6.2) A formal nutrition program is recommended to meet the behavioral and nutritional needs of all 

species and specimens within the collection.



(2.6.3) Animal diets must be of a quality and quantity suitable for each animal’s nutritional and 

psychological needs. Diet formulations and records of analysis of appropriate feed items should be 

maintained and may be examined by the Visiting Committee. Animal food, especially seafood 

products, should be purchased from reliable sources that are sustainable and/or well managed. 

(2.6.1) Animal food preparations must meet all local, state/provincial, and federal regulations. 

(2.6.4) The institution should assign at least one person to oversee appropriate browse material for the 

collection. 

Chapter 6 

(2.1.1) A full-time staff veterinarian is recommended. However, the Commission realizes that in some 

cases such is not practical. In those cases, a consulting/part-time veterinarian must be under contract 

to make at least twice monthly inspections of the animal collection and respond as soon as possible 

to any emergencies. The Commission also recognizes that certain collections, because of their size 

and/or nature, may require different considerations in veterinary care. 

(2.1.2) So that indications of disease, injury, or stress may be dealt with promptly, veterinary coverage 

must be available to the animal collection 24 hours a day, 7 days a week. 

(2.2.1) Written, formal procedures must be available to the animal care staff for the use of animal drugs 

for veterinary purposes and appropriate security of the drugs must be provided. 

(1.4.6) A staff member must be designated as being responsible for the institution's animal recordkeeping 

system. That person must be charged with establishing and maintaining the institution's 

animal records, as well as with keeping all animal care staff members apprised of relevant laws and 

regulations regarding the institution's animal collection. 

(1.4.7) Animal records must be kept current, and data must be logged daily. 

(1.4.5) At least one set of the institution’s historical animal records must be stored and protected. Those 

records should include permits, titles, declaration forms, and other pertinent information. 

(1.4.4) Animal records, whether in electronic or paper form, including health records, must be duplicated 

and stored in a separate location. 

(1.4.3) Animals must be identifiable, whenever practical, and have corresponding ID numbers. For 

animals maintained in colonies or other animals not considered readily identifiable, the institution 

must provide a statement explaining how record keeping is maintained. 

(1.4.1) An animal inventory must be compiled at least once a year and include data regarding acquisitions 

and dispositions in the animal collection. 

(1.4.2) All species owned by the institution must be listed on the inventory, including those animals on 

loan to and from the institution. In both cases, notations should be made on the inventory. 

(2.7.1) The institution must have holding facilities or procedures for the quarantine of newly arrived 

animals and isolation facilities or procedures for the treatment of sick/injured animals. 

(2.7.3) Quarantine, hospital, and isolation areas should be in compliance with standards or guidelines 

adopted by the AZA. 

(2.7.2) Written, formal procedures for quarantine must be available and familiar to all staff working with 

quarantined animals. 

(11.1.2) Training and procedures must be in place regarding zoonotic diseases. 

(11.1.3) A tuberculin testing and surveillance program must be established for appropriate staff in order to 

ensure the health of both the employees and the animal collection. 

(2.5.1) Deceased animals should be necropsied to determine the cause of death. Disposal after necropsy 

must be done in accordance with local/federal laws. 

(2.4.1) The veterinary care program must emphasize disease prevention. 

(1.5.5) For animals used in offsite programs and for educational purposes, the institution must have 

adequate protocols in place to protect the rest of the collection from exposure to infectious agents. 

(2.3.1) Capture equipment must be in good working order and available to authorized, trained personnel 

at all times. 

(2.4.2) Keepers should be trained to recognize abnormal behavior and clinical symptoms of illness and 

have knowledge of the diets, husbandry (including enrichment items and strategies), and restraint 

procedures required for the animals under their care. However, keepers should not evaluate illnesses 

nor prescribe treatment. 

(2.3.2) Hospital facilities should have x-ray equipment or have access to x-ray services.



(1.5.8) The institution must develop a clear process for identifying and addressing animal welfare 

concerns within the institution. 

Chapter 8 

(1.6.1) The institution must have a formal written enrichment program that promotes species-appropriate 

behavioral opportunities. 

(1.6.2) The institution must have a specific staff member(s) or committee assigned for enrichment 

program oversight, implementation, training, and interdepartmental coordination of enrichment efforts. 

Chapter 9 

(5.3) A written policy on the use of live animals in programs should be on file. Animals in education 

programs must be maintained and cared for by trained staff, and housing conditions must meet 

standards set for the remainder of the animal collection, including species-appropriate shelter, 

exercise, social and environmental enrichment, access to veterinary care, nutrition, etc. Since some 

of these requirements can be met outside of the primary enclosure, for example, enclosures may be 

reduced in size provided that the animal’s physical and psychological needs are being met. 

(1.5.3) If animal demonstrations are a part of the institution’s programs, an education and conservation 

message must be an integral component. 

Chapter 10 

(5.3) Institutions should maximize the generation of scientific knowledge gained from the animal 

collection. This might be achieved by participating in AZA TAG/SSP sponsored research when 

applicable, conducting original research projects, affiliating with local universities, and/or employing 

staff with scientific credentials. 

(5.2) Institutions must have a written policy that outlines the type of research that it conducts, methods, 

staff involvement, evaluations, animals to be involved, and guidelines for publication of findings. 

(5.1) Research activities must be under the direction of a person qualified to make informed decisions 

regarding research.



Appendix B: Acquisition/Disposition Policy 

I. Introduction: The Association of Zoos and Aquariums (AZA) was established, among other reasons, to 

foster continued improvement in the zoological park and aquarium profession. One of its most important 

roles is to provide a forum for debate and consensus building among its members, the intent of which is 

to attain high ethical standards, especially those related to animal care and professional conduct. The 

stringent requirements for AZA accreditation and high standards of professional conduct are unmatched 

by similar organizations and also far surpass the United States Department of Agriculture's Animal and 

Plant Health Inspection Service's requirements for licensed animal exhibitors. AZA member facilities must 

abide by a Code of Professional Ethics - a set of standards that guide all aspects of animal management 

and welfare. As a matter of priority, AZA institutions should acquire animals from other AZA institutions 

and dispose of animals to other AZA institutions. 

AZA accredited zoological parks and aquariums cannot fulfill their important missions of conservation, 

education and science without living animals. Responsible management of living animal populations 

necessitates that some individuals be acquired and that others be removed from the collection at certain 

times. Acquisition of animals can occur through propagation, trade, donation, loan, purchase, capture, or 

rescue. Animals used as animal feed are not accessioned into the collection. 

Disposition occurs when an animal leaves the collection for any reason. Reasons for disposition vary 

widely, but include cooperative population management (genetic or demographic management), 

reintroduction, behavioral incompatibility, sexual maturation, animal health concerns, loan or transfer, or 

death. 

The AZA Acquisition/Disposition Policy (A/D) was created to help (1) guide and support member 

institutions in their animal acquisition and disposition decisions, and (2) ensure that all additions and 

removals are compatible with the Association's stated commitment to "save and protect the wonders of 

the living natural world." More specifically, the AZA A/D Policy is intended to: 

• Ensure that the welfare of individual animals and conservation of populations, species and 

ecosystems are carefully considered during acquisition and disposition activities; 

• Maintain a proper standard of conduct for AZA members during acquisition and disposition 

activities; 

• Ensure that animals from AZA member institutions are not transferred to individuals or 

organizations that lack the appropriate expertise or facilities to care for them. 

• Support the goal of AZA’s cooperatively managed populations and associated programs, 

including Species Survival Plans® (SSPs), Population Management Plans (PMPs), and Taxon 

Advisory Groups (TAGs). 

The AZA Acquisition/Disposition Policy will serve as the default policy for AZA member institutions. 

Institutions may develop their own A/D Policy in order to address specific local concerns. Any institutional 

policy must incorporate and not conflict with the AZA acquisition and disposition standards. 

Violations of the AZA Acquisition/Disposition Policy will be dealt with in accordance with the AZA 

Code of Professional Ethics. Violations can result in an institution's or individual's expulsion from 

membership in the AZA. 

II. Group or Colony-based Identification: For some colonial, group-living, or prolific species, such as 

certain insects, aquatic invertebrates, schooling fish, rodents, and bats, it is often impossible or highly 

impractical to identify individual specimens. These species are therefore maintained, acquisitioned, and 

disposed of as a group or colony. Therefore, when this A/D Policy refers to animals or specimens, it is in 

reference to both individuals and groups/colonies. 

III. Germplasm: Acquisition and disposition of germplasm should follow the same guidelines outlined in 

this document if its intended use is to create live animal(s). Ownership of germplasm and any resulting 

animals should be clearly defined. Institutions acquiring or dispositioning germplasm or any animal parts 

or samples should consider not only its current use, but also future possible uses as new technologies 

become available. 

IV(a). General Acquisitions: Animals are to be acquisitioned into an AZA member institution’s collection 

if the following conditions are met:



1. Acquisitions must meet the requirements of all applicable local, state, federal and international 

regulations and laws. 

2. The Director or Chief Executive Officer of the institution is charged with the final authority and 

responsibility for the monitoring and implementation of all acquisitions. 

3. Acquisitions must be consistent with the mission of the institution, as reflected in its Institutional 

Collection Plan, by addressing its exhibition/education, conservation, and/or scientific goals. 

4. Animals that are acquired for the collection, permanently or temporarily, must be listed on 

institutional records. All records should follow the Standards for Data Entry and Maintenance of 

North American Zoo and Aquarium Animal Records Databases ® . 

5. Animals may be acquired temporarily for reasons such as, holding for governmental agencies, 

rescue and/or rehabilitation, or special exhibits. Animals should only be accepted if they will not 

jeopardize the health, care or maintenance of the animals in the permanent collection or the animal 

being acquired. 

6. The institution must have the necessary resources to support and provide for the professional care 

and management of a species, so that the physical and social needs of both specimen and species 

are met. 

7. Attempts by members to circumvent AZA conservation programs in the acquisition of SSP animals 

are detrimental to the Association and its conservation programs. Such action may be detrimental 

to the species involved and is a violation of the Association's Code of Professional Ethics. All AZA 

members must work through the SSP program in efforts to acquire SSP species and adhere to the 

AZA Full Participation policy. 

8. Animals are only to be acquired from sources that are known to operate legally and conduct their 

business in a manner that reflects and/or supports the spirit and intent of the AZA Code of 

Professional Ethics as well as this policy. Any convictions of state, federal, or international wildlife 

laws should be reviewed, as well as any previous dealings with other AZA accredited institutions. 

9. When acquiring specimens managed by a PMP, institutions should consult with the PMP manager. 

10. Institutions should consult AZA Wildlife Conservation and Management Committee (WCMC)approved 

Regional Collection Plans (RCPs) when making acquisition decisions. 

IV(b). Acquisitions from the Wild: The maintenance of wild animal populations for education and wildlife 

conservation purposes is a unique responsibility of AZA member zoos and aquariums. To accomplish 

these goals, it may be necessary to acquire wild-caught specimens. Before acquiring animals from the 

wild, institutions are encouraged to examine sources including other AZA institutions or regional 

zoological associations. 

When acquiring animals from the wild, careful consideration must be taken to evaluate the long-term 

impacts on the wild population. Any capture of free-ranging animals should be done in accordance with all 

local, state, federal, and international wildlife laws and regulations and not be detrimental to the long-term 

viability of the species or the wild or ex-situ population(s). In crisis situations, when the survival of a 

population is at risk, rescue decisions are to be made on a case-by-case basis. 

V(a). Disposition Requirements – Living Animals: Successful conservation and animal management 

efforts rely on the cooperation of many entities, both within and outside of AZA. While preference is given 

to placing animals within AZA member institutions, it is important to foster a cooperative culture among 

those who share the primary mission of AZA accredited facilities. The AZA draws a strong distinction 

between the mission, stated or otherwise, of non-AZA member organizations and the mission of 

professionally managed zoological parks and aquariums accredited by the AZA. 

An accredited AZA member balances public display, recreation, and entertainment with demonstrated 

efforts in education, conservation, and science. While some non-AZA member organizations may meet 

minimum daily standards of animal care for wildlife, the AZA recognizes that this, by itself, is insufficient to 

warrant either AZA membership or participation in AZA's cooperative animal management programs. 

When an animal is sent to a non-member of AZA, it is imperative that the member be confident that the 

animal will be cared for properly. 

Animals may only be disposed of from an AZA member institution's collection if the following 

conditions are met:



1. Dispositions must meet the requirements of all applicable local, state, federal and international 

regulations and laws. 

2. The Director or Chief Executive Officer of the institution is charged with the final authority and 

responsibility for the monitoring and implementation of all dispositions. 

3. Any disposition must abide by the Mandatory Standards and General Advisories of the AZA Code 

of Professional Ethics. Specifically, "a member shall make every effort to assure that all animals in 

his/her collection and under his/her care are disposed of in a manner which meets the current 

disposition standards of the Association and do not find their way into the hands of those not 

qualified to care for them properly." 

4. Non-domesticated animals shall not be disposed of at animal auctions. Additionally, animals shall 

not be disposed of to any organization or individual that may use or sell the animal at an animal 

auction. In transactions with AZA non-members, the recipient must ensure in writing that neither the 

animal nor its offspring will be disposed of at a wild animal auction or to an individual or 

organization that allows the hunting of the animal. 

5. Animals shall not be disposed of to organizations or individuals that allow the hunting of these 

animals or their offspring. This does not apply to individuals or organizations which allow the 

hunting of only free-ranging game species (indigenous to North America) and established longintroduced 

species such as, but not limited to, white-tailed deer, quail, rabbit, waterfowl, boar, ringnecked 

pheasant, chukar, partridge, and trout. AZA distinguishes hunting/fishing for sport from 

culling for sustainable population management and wildlife conservation purposes. 

6. Attempts by members to circumvent AZA conservation programs in the disposition of SSP animals 

are detrimental to the Association and its conservation programs. Such action may be detrimental 

to the species involved and is a violation of the Association's Code of Professional Ethics. All AZA 

members must work through the SSP program in efforts to deacquisition SSP species and adhere 

to the AZA Full Participation policy. 

7. Domesticated animals are to be disposed of in a manner consistent with acceptable farm practices 

and subject to all relevant laws and regulations. 

8. Live specimens may be released within native ranges, subject to all relevant laws and regulations. 

Releases may be a part of a recovery program and any release must be compatible with the AZA 

Guidelines for Reintroduction of Animals Born or Held in Captivity, dated June 3, 1992. 

9. Detailed disposition records of all living or dead specimens must be maintained. Where applicable, 

proper animal identification techniques should be utilized. 

10. It is the obligation of every loaning institution to monitor, at least annually, the conditions of any 

loaned specimens and the ability of the recipient to provide proper care. If the conditions and care 

of animals are in violation of the loan agreement, it is the obligation of the loaning institution to 

recall the animal. Furthermore, an institution's loaning policy must not be in conflict with this A/D 

Policy. 

11. If live specimens are euthanized, it must be done in accordance with the established policy of the 

institution and the Report of the American Veterinary Medical Association Panel on Euthanasia 

(Journal of the American Veterinary Medical Association 218 (5): 669-696, 2001). 

12. In dispositions to non-AZA members, the non-AZA member's mission (stated or implied) must not 

be in conflict with the mission of AZA, or with this A/D Policy. 

13. In dispositions to non-AZA member facilities that are open to the public, the non-AZA member must 

balance public display, recreation, and entertainment with demonstrated efforts in conservation, 

education, and science. 

14. In dispositions to non-AZA members, the AZA members must be convinced that the recipient has 

the expertise, records management practices, financial stability, facilities, and resources required to 

properly care for and maintain the animals and their offspring. It is recommended that this 

documentation be kept in the permanent record of the animals at the AZA member institution. 

15. If living animals are sent to a non-AZA member research institution, the institution must be 

registered under the Animal Welfare Act by the U.S. Department of Agriculture Animal and Plant 

Health Inspection Service. For international transactions, the receiving facility should be registered 

by that country's equivalent body with enforcement over animal welfare. 

16. No animal disposition should occur if it would create a health or safety risk (to the animal or 

humans) or have a negative impact on the conservation of the species.



17. Inherently dangerous wild animals or invasive species should not be dispositioned to the pet trade 

or those unqualified to care for them. 

18. Under no circumstances should any primates be dispositioned to a private individual or to the pet 

trade. 

19. Fish and aquatic invertebrate species that meet ANY of the following are inappropriate to be 

disposed of to private individuals or the pet trade: 

a. species that grow too large to be housed in a 72-inch long, 180 gallon aquarium (the largest tank 

commonly sold in retail stores) 

b. species that require extraordinary life support equipment to maintain an appropriate ex-situ 

environment (e.g., cold water fish and invertebrates) 

c. species deemed invasive (e.g., snakeheads) 

d. species capable of inflicting a serious bite or venomous sting (e.g., piranha, lion fish, blue-ringed 

octopus) 

e. species of wildlife conservation concern 

20. When dispositioning specimens managed by a PMP, institutions should consult with the PMP 

manager. 

21. Institutions should consult WCMC-approved RCPs when making disposition decisions. 

V(b). Disposition Requirements – Dead Specimens: Dead specimens (including animal parts and 

samples) are only to be disposed of from an AZA member institution's collection if the following conditions 

are met: 

1. Dispositions of dead specimens must meet the requirements of all applicable local, state, federal 

and international regulations and laws. 

2. Maximum utilization is to be made of the remains, which could include use in educational programs 

or exhibits. 

3. Consideration is given to scientific projects that provide data for species management and/or 

conservation. 

4. Records (including ownership information) are to be kept on all dispositions, including animal body 

parts, when possible. 

5. SSP and TAG necropsy protocols are to be accommodated insofar as possible. 

VI. Transaction Forms: AZA member institutions will develop transaction forms to record animal 

acquisitions and dispositions. These forms will require the potential recipient or provider to adhere to the 

AZA Code of Professional Ethics, the AZA Acquisition/Disposition Policy, and all relevant AZA and 

member policies, procedures and guidelines. In addition, transaction forms must insist on compliance with 

the applicable laws and regulations of local, state, federal and international authorities.



Appendix C: Recommended Quarantine Procedures 

Quarantine Facility: A separate quarantine facility, with the ability to accommodate mammals, birds, 

reptiles, amphibians, and fish should exist. If a specific quarantine facility is not present, then newly 

acquired animals should be isolated from the established collection in such a manner as to prohibit 

physical contact, to prevent disease transmission, and to avoid aerosol and drainage contamination. 

Such separation should be obligatory for primates, small mammals, birds, and reptiles, and attempted 

wherever possible with larger mammals such as large ungulates and carnivores, marine mammals, and 

cetaceans. If the receiving institution lacks appropriate facilities for isolation of large primates, preshipment 

quarantine at an AZA or AALAS accredited institution may be applied to the receiving 

institutions protocol. In such a case, shipment must take place in isolation from other primates. More 

stringent local, state, or federal regulations take precedence over these recommendations. 

Quarantine Length: Quarantine for all species should be under the supervision of a veterinarian and 

consist of a minimum of 30 days (unless otherwise directed by the staff veterinarian). Mammals: If during 

the 30-day quarantine period, additional mammals of the same order are introduced into a designated 

quarantine area, the 30-day period must begin over again. However, the addition of mammals of a 

different order to those already in quarantine will not have an adverse impact on the originally quarantined 

mammals. Birds, Reptiles, Amphibians, or Fish: The 30-day quarantine period must be closed for each of 

the above Classes. Therefore, the addition of any new birds into a bird quarantine area requires that the 

30-day quarantine period begin again on the date of the addition of the new birds. The same applies for 

reptiles, amphibians, or fish. 

Quarantine Personnel: A keeper should be designated to care only for quarantined animals or a keeper 

should attend quarantined animals only after fulfilling responsibilities for resident species. Equipment 

used to feed and clean animals in quarantine should be used only with these animals. If this is not 

possible, then equipment must be cleaned with an appropriate disinfectant (as designated by the 

veterinarian supervising quarantine) before use with post-quarantine animals. 

Institutions must take precautions to minimize the risk of exposure of animal care personnel to 

zoonotic diseases that may be present in newly acquired animals. These precautions should include the 

use of disinfectant foot baths, wearing of appropriate protective clothing and masks in some cases, and 

minimizing physical exposure in some species; e.g., primates, by the use of chemical rather than physical 

restraint. A tuberculin testing/surveillance program must be established for zoo/aquarium employees in 

order to ensure the health of both the employees and the animal collection. 

Quarantine Protocol: During this period, certain prophylactic measures should be instituted. Individual 

fecal samples or representative samples from large numbers of individuals housed in a limited area (e.g., 

birds of the same species in an aviary or frogs in a terrarium) should be collected at least twice and 

examined for gastrointestinal parasites. Treatment should be prescribed by the attending veterinarian. 

Ideally, release from quarantine should be dependent on obtaining two negative fecal results spaced a 

minimum of two weeks apart either initially or after parasiticide treatment. In addition, all animals should 

be evaluated for ectoparasites and treated accordingly. 

Vaccinations should be updated as appropriate for each species. If the animal arrives without a 

vaccination history, it should be treated as an immunologically naive animal and given an appropriate 

series of vaccinations. Whenever possible, blood should be collected and sera banked. Either a -94°F 

(-70ºC) frost-free freezer or a -4°F (-20ºC) freezer that is not frost-free should be available to save sera. 

Such sera could provide an important resource for retrospective disease evaluation. 

The quarantine period also represents an opportunity to, where possible, permanently identify all 

unmarked animals when anesthetized or restrained (e.g., tattoo, ear notch, ear tag, etc.). Also, whenever 

animals are restrained or immobilized, a complete physical, including a dental examination, should be 

performed. Complete medical records should be maintained and available for all animals during the 

quarantine period. Animals that die during quarantine should have a necropsy performed under the 

supervision of a veterinarian and representative tissues submitted for histopathologic examination. 

Quarantine Procedures: The following are recommendations and suggestions for appropriate 

quarantine procedures for Lutrinae:

Required: 

1. Direct and flotation fecals 

2. Vaccinate as appropriate 

Strongly Recommended: 

1. CBC/sera profile 

2. Urinalysis 

3. Appropriate serology (FIP, FeLV, FIV) 

4. Heartworm testing in appropriate species 


Otter (Lutrinae) Care Manual



Appendix D: Program Animal Position Statement 

The Conservation Education Committee (CEC) of the Association of Zoos and Aquariums supports 

the appropriate use of program animals as an important and powerful educational tool that provides a 

variety of benefits to zoo and aquarium educators seeking to convey cognitive and affective (emotional) 

messages about conservation and wildlife. Utilizing these animals allows educators to strongly engage 

audiences. As discussed below, the use of program animals has been demonstrated to result in 

lengthened learning periods, increased knowledge acquisition and retention, enhanced environmental 

attitudes, and the creation of positive perceptions concerning zoo and aquarium animals. 

Audience Engagement: Zoos and aquariums are ideal venues for developing emotional ties to wildlife 

and fostering an appreciation for the natural world. However, developing and delivering effective 

educational messages in the free-choice learning environments of zoos and aquariums is a difficult task. 

Zoo and aquarium educators are constantly challenged to develop methods for engaging and teaching 

visitors who often view a trip to the zoo as a social or recreational experience (Morgan & Hodgkinson 

1999). The use of program animals can provide the compelling experience necessary to attract and 

maintain personal connections with visitors of all motivations, thus preparing them for learning and 

reflection on their own relationships with nature. 

Program animals are powerful catalysts for learning for a variety of reasons. They are generally 

active, easily viewed, and usually presented in close proximity to the public. These factors have proven to 

contribute to increasing the length of time that people spend watching animals in zoo exhibits (Wolf & 

Tymitz 1981; Bitgood, Patterson & Benefield, 1986, 1988). In addition, the provocative nature of a 

handled animal likely plays an important role in captivating a visitor. In two studies (Povey & Rios 2002; 

Povey 2002), visitors viewed animals three and four times longer while they were being presented in 

demonstrations outside of their enclosure with an educator than while they were on exhibit. Clearly, the 

use of program animals in shows or informal presentations is effective in lengthening the potential time 

period for learning and overall impact. 

Program animals also provide the opportunity to personalize the learning experience, tailoring the 

teaching session to what interests the visitors. Traditional graphics offer little opportunity for this level of 

personalization of information delivery and are frequently not read by visitors (Churchman 1985; Johnston 

1998). For example, Povey (2002) found that only 25% of visitors to an animal exhibit read the 

accompanying graphic; whereas, 45% of visitors watching the same animal handled in an educational 

presentation asked at least one question and some asked as many as seven questions. Having an animal 

accompany the educator allowed the visitors to make specific inquiries about topics in which they were 

interested. 

Knowledge Acquisition: Improving our visitors' knowledge and understanding regarding wildlife and 

wildlife conservation is a fundamental goal for many zoo educators using program animals. A growing 

body of evidence supports the validity of using program animals to enhance delivery of these cognitive 

messages as well. 

• MacMillen (1994) found that the use of live animals in a zoomobile outreach program significantly 

enhanced cognitive learning in a vertebrate classification unit for sixth grade students. 

• Sherwood et al. (1989) compared the use of live horseshoe crabs and sea stars to the use of 

dried specimens in an aquarium education program and demonstrated that students made the 

greatest cognitive gains when exposed to programs utilizing the live animals. 

• Povey and Rios (2002) noted that in response to an open-ended survey question (“Before I saw 

this animal, I never realized that…”), visitors watching a presentation utilizing a program animal 

provided 69% cognitive responses (i.e., something they learned) versus 9% made by visitors 

viewing the same animal in its exhibit (who primarily responded with observations). 

• Povey (2002) recorded a marked difference in learning between visitors observing animals on 

exhibit versus being handled during informal presentations. Visitors to demonstrations utilizing a 

raven and radiated tortoises were able to answer questions correctly at a rate as much as eleven 

times higher than visitors to the exhibits. 

Enhanced Environmental Attitudes: Program animals have been clearly demonstrated to increase 

effective learning and attitudinal change.



• Studies by Yerke and Burns (1991) and Davison et al. (1993) evaluated the effect live animal 

shows had on visitor attitudes. Both found their shows successfully influenced attitudes about 

conservation and stewardship. 

• Yerke and Burns (1993) also evaluated a live bird outreach program presented to Oregon fifthgraders 

and recorded a significant increase in students' environmental attitudes after the 

presentations. 

• Sherwood et al. (1989) found that students who handled live invertebrates in an education 

program demonstrated both short and long-term attitudinal changes as compared to those who 

only had exposure to dried specimens. 

• Povey and Rios (2002) examined the role program animals play in helping visitors develop 

positive feelings about the care and well-being of zoo animals. 

• As observed by Wolf and Tymitz (1981), zoo visitors are deeply concerned with the welfare of zoo 

animals and desire evidence that they receive personalized care. 

Conclusion: Creating positive impressions of aquarium and zoo animals, and wildlife in general, is 

crucial to the fundamental mission of zoological institutions. Although additional research will help us 

delve further into this area, the existing research supports the conclusion that program animals are an 

important tool for conveying both cognitive and affective messages regarding animals and the need to 

conserve wildlife and wild places.



Appendix E: Developing an Institutional Program Animal Policy 

Membership in AZA requires that an institution meet the AZA Accreditation Standards collectively 

developed by our professional colleagues. Standards guide all aspects of an institution's operations; 

however, the accreditation commission has asserted that ensuring that member institutions demonstrate 

the highest standards of animal care is a top priority. Another fundamental AZA criterion for membership 

is that education be affirmed as core to an institution’s mission. All accredited public institutions are 

expected to develop a written education plan and to regularly evaluate program effectiveness. 

The inclusion of animals (native, exotic and domestic) in educational presentations, when done 

correctly, is a powerful tool. CEC's Program Animal Position Statement (Appendix D) describes the 

research underpinning the appropriate use of program animals as an important and powerful educational 

tool that provides a variety of benefits to zoo and aquarium educators seeking to convey cognitive and 

affective messages about conservation and wildlife. Ongoing research, such as AZA's Multi-Institutional 

Research Project (MIRP) and research conducted by individual AZA institutions will help zoo educators to 

determine whether the use of program animals conveys intended and conflicting messages and to modify 

and improve programs accordingly. 

When utilizing program animals our responsibility is to meet both our high standards of animal care 

and our educational goals. Additionally, as animal management professionals, we must critically address 

both the species' conservation needs and the welfare of the individual animal. Because "wild creatures 

differ endlessly," in their forms, needs, behavior, limitations and abilities (Conway 1995), AZA, through its 

Animal Welfare Committee, has recently given the responsibility to develop taxon-specific animal welfare 

standards to the Taxon Advisory Groups (TAG) and Species Survival Plan ® Program (SSP). Experts 

within each TAG or SSP, along with their education advisors, are charged with assessing all aspects of 

the taxons’ biological and social needs and developing animal care standards that include specifications 

concerning their use as program animals. 

However, even the most exacting standards cannot address the individual choices faced by each 

AZA institution. Therefore, each institution is required to develop a program animal policy that articulates 

and evaluates program benefits. The following recommendations are offered to assist each institution in 

formulating its own Institutional Program Animal Policy. 

The Policy Development Process: Within each institution, key stakeholders should be included in the 

development of that institution's policy, including, but not limited to representatives from: 

• The Education Department 

• The Animal Husbandry Department 

• The Veterinary and Animal Health Department 

• The Conservation & Science Department 

• Any animal show staff (if in a separate department) 

• Departments that frequently request special program animal situations (e.g., special events, 

development, marketing, zoo or aquarium society, administration) 

• Additionally, staff from all levels of the organization should be involved in this development (e.g., 

curators, keepers, education managers, interpreters, volunteer coordinators). 

To develop a comprehensive Program Animal Policy, we recommend that the following components 

be included: 

I. Philosophy: In general, the position of the AZA is that the use of animals in up close and personal 

settings, including animal contact, can be extremely positive and powerful, as long as: 

• The use and setting is appropriate. 

• Animal and human welfare is considered at all times. 

• The animal is used in a respectful, safe manner and in a manner that does not misrepresent or 

degrade the animal. 

• A meaningful conservation message is an integral component. Read the AZA Board-approved 

Conservation Messages. 

• Suitable species and individual specimens are used. 

Institutional program animal policies should include a philosophical statement addressing the above, 

and should relate the use of program animals to the institution's overall mission statement.



II. Appropriate Settings: The Program Animal Policy should include a listing of all settings both on and 

off site, where program animal use is permitted. This will clearly vary among institutions. Each institution's 

policy should include a comprehensive list of settings specific to that institution. Some institutions may 

have separate policies for each setting; others may address the various settings within the same policy. 

Examples of settings include: 

On-site programming: 

Informal and non-registrants: 

• On-grounds programming with animals being brought out (demonstrations, lectures, parties, 

special events, and media) 

• Children's zoos and contact yards 

• Behind-the-scenes open houses 

• Shows 

• Touch pools 

Formal (registration involved) and controlled settings: 

• School group programs 

• Summer Camps 

• Overnights 

• Birthday parties 

Offsite and outreach: 

• PR events (TV, radio) 

• Fundraising events 

• Field programs involving the public 

• School visits 

• Library visits 

• Nursing Home visits (therapy) 

• Hospital visits 

• Senior Centers 

• Civic Group events 

In some cases, policies will differ from setting to setting (e.g., on-site and off-site use with media). 

These settings should be addressed separately, and should reflect specific animal health issues, 

assessment of stress in these situations, limitations, and restrictions. 

III. Compliance with Regulations: All AZA institutions housing mammals are regulated by the USDA’s 

Animal Welfare Act. Other federal regulations, such as the Marine Mammal Protection Act, may apply. 

Additionally, many states, and some cities, have regulations that apply to animal contact situations. 

Similarly, all accredited institutions are bound by the AZA Code of Professional Ethics. It is expected that 

the Institution Program Animal Policy address compliance with appropriate regulations and AZA 

Accreditation Standards. 

IV. Collection Planning: All AZA accredited institutions should have a collection planning process in 

place. Program animals are part of an institution's overall collection and must be included in the overall 

collection planning process. The AZA Guide to Accreditation contains specific requirements for the 

institution collection plan. For more information about collection planning in general, please see the 

Collection Management pages in the Members Only section of the AZA website (www.aza.org). The 

following recommendations apply to program animals: 

1. Listing of approved program animals (to be periodically amended as collection changes). 

Justification of each species should be based upon criteria such as: 

a. Temperament and suitability for program use 

b. Husbandry requirements 

c. Husbandry expertise 

d. Veterinary issues and concerns 

e. Ease and means of acquisition / disposition 

f. Educational value and intended conservation message 

g. Conservation Status 

h. Compliance with TAG and SSP guidelines and policies



2. General guidelines as to how each species (and, where necessary, for each individual) will be 

presented to the public, and in what settings 

3. The collection planning section should reference the institution's acquisition and disposition 

policies. 

V. Conservation Education Message: As noted in the AZA Accreditation Standards, if animal 

demonstrations are part of an institution's programs, an educational and conservation message must be 

an integral component. The Program Animal Policy should address the specific messages related to the 

use of program animals, as well as the need to be cautious about hidden or conflicting messages (e.g., 

“petting” an animal while stating verbally that it makes a poor pet). This section may include or reference 

the AZA Conservation Messages. 

Although education value and messages should be part of the general collection planning process, 

this aspect is so critical to the use of program animals that it deserves additional attention. In addition, it is 

highly recommended to encourage the use of biofacts in addition to or in place of the live animals. 

Whenever possible, evaluation of the effectiveness of presenting program animals should be built into 

education programs. 

VI. Human Health and Safety: The safety of our staff and the public is one of the greatest concerns in 

working with program animals. Although extremely valuable as educational and affective experiences, 

contact with animals poses certain risks to the handler and the public. Therefore, the human health and 

safety section of the policy should address: 

• Minimization of the possibility of disease transfer from non-human animals to humans, and viceversa 

(e.g., hand washing stations, no touch policies, use of hand sanitizer). 

• Safety issues related to handlers' personal attire and behavior (e.g., discourage or prohibit use of 

long earrings, perfume and cologne, not eating or drinking around animals, smoking etc.). 

AZA’s Animal Contact Policy provides guidelines in this area; these guidelines were incorporated into 

accreditation standards in 1998. 

VII. Animal Health and Welfare: Animal health and welfare are the highest priority of AZA accredited 

institutions. As a result, the Institutional Program Animal Policy should make a strong statement on the 

importance of animal welfare. The policy should address: 

• General housing, husbandry, and animal health concerns (e.g. that the housing and husbandry 

for program animals meets or exceeds general standards and that the needs of the individual 

animal, such as enrichment and visual cover, are accommodated). 

• The empowerment of handlers to make decisions related to animal health and welfare; such as 

withdrawing animals from a situation if safety or health is in danger of being compromised. 

• Requirements for supervision of contact areas and touch tanks by trained staff and volunteers. 

• Frequent evaluation of human/animal interactions to assess safety, health, welfare, etc. 

• Ensure that the level of health care for the program animals is consistent with that of other 

animals in the collection. 

VIII. Taxon Specific Protocols: The AZA encourages institutions to provide taxonomically specific 

protocols, either at the genus or species level, or the specimen, or individual, level. Some taxon-specific 

guidelines may affect the use of program animals. To develop these, institutions refer to the Conservation 

Programs Database. Taxon-specific protocols should address: 

• How to remove the individual animal from and return it to its permanent enclosure. 

• How to crate and transport animals. 

• Signs of stress, stress factors and discomfort behaviors. 

• Situation specific handling protocols (e.g., whether or not animal is allowed to be touched by the 

public, and how to handle in such situations) 

• Guidelines for disinfecting surfaces, transport carriers, enclosures, etc. 

• Animal facts and conservation information. 

• Limitations and restrictions regarding ambient temperatures and or weather conditions. 

• Time limitations (including animal rotation and rest periods, as appropriate, duration of time each 

animal can participate, and restrictions on travel distances).



• The numbers of trained personnel required to ensure the health and welfare of the animals, 

handlers and public. 

• Taxon-specific guidelines on animal health. 

IX. Logistics, and Managing the Program: The Institutional Policy should address a number of logistical 

issues related to program animals, including: 

• Where and how the program animal collection will be housed, including any quarantine and 

separation for animals used off-site. 

• Procedures for requesting animals, including the approval process and decision making process. 

• Accurate documentation and availability of records, including procedures for documenting animal 

usage, animal behavior, and any other concerns that arise. 

X. Staff Training: Thorough training for all handling staff (keepers, educators, and volunteers, and 

docents) is clearly critical. Staff training is such a large issue that many institutions may have separate 

training protocols and procedures. Specific training protocols can be included in the Institutional Program 

Animal Policy or reference can be made that a separate training protocol exists. It is recommended that 

the training section of the policy address: 

• Personnel authorized to handle and present animals. 

• Handling protocol during quarantine. 

• The process for training, qualifying and assessing handlers including who is authorized to train 

handlers. 

• The frequency of required re-training sessions for handlers. 

• Personnel authorized to train animals and training protocols. 

• The process for addressing substandard performance and noncompliance with established 

procedures. 

• Medical testing and vaccinations required for handlers (e.g., TB testing, tetanus shots, rabies 

vaccinations, routine fecal cultures, physical exams, etc.). 

• Training content (e.g., taxonomically specific protocols, natural history, relevant conservation 

education messages, presentation techniques, interpretive techniques). 

• Protocols to reduce disease transmission (e.g., zoonotic disease transmission, proper hygiene 

and hand washing requirements, as noted in AZA's Animal Contact Policy). 

• Procedures for reporting injuries to the animals, handling personnel or public. 

• Visitor management (e.g., ensuring visitors' interact appropriately with animals, do not eat or drink 

around the animal, etc.). 

XI. Review of Institutional Policies: All policies should be reviewed regularly. Accountability and 

ramifications of policy violations should be addressed as well (e.g., retraining, revocation of handling 

privileges, etc.). Institutional policies should address how frequently the Program Animal Policy will be 

reviewed and revised, and how accountability will be maintained. 

XII. TAG and SSP Recommendations: Following development of taxon-specific recommendations from 

each TAG and SSP, the institution policy should include a statement regarding compliance with these 

recommendations. If the institution chooses not to follow these specific recommendations, a brief 

statement providing rationale is recommended.



Appendix F: AZA Accreditation Standards for Otters 

The following specific standards of care and recommendations for otters are taken from the AZA 

Accreditation Standards and Related Policies (AZA 2008): 

Water Quality: The provision of fresh potable water is a requirement of USDA Animal Welfare 

Regulations (AWR 2005) as stated: “If potable water is not accessible to the animals at all times, it must 

be provided as often as necessary for the health and comfort of the animal. Frequency of watering shall 

consider age, species, condition, size, and type of the animal. All water receptacles shall be kept clean 

and sanitary” (AWR 2005). Considering the needs of otters, the AZA Small Carnivore TAG state that 

otters should be given fresh water daily if their pools are not filtered or dumped and filled daily on a daily 

basis. AZA Accreditation Standards require that institutions abide by relevant federal laws and 

regulations: 

“The institution must comply with all relevant local, state, and federal wildlife laws and regulations. It is 

understood that, in some cases, AZA accreditation standards are more stringent than existing laws and 

regulations. In these cases the AZA standard must be met” (AZA 2008). 

Transport: The standards of care identified in Chapter 3 are based on IATA regulations (IATA 2007). 

Institutions transporting otters are obliged to abide by these regulations as stated in the following AZA 

Accreditation Standard: 

“The institution must comply with all relevant local, state, and federal wildlife laws and regulations. It is 

understood that, in some cases, AZA accreditation standards are more stringent than existing laws and 

regulations. In these cases the AZA standard must be met” (AZA 2008). 

Quarantine: “Quarantine, hospital, and isolation areas should be in compliance with standards/guidelines 

adopted by the AZA” (AZA 2008) 

Quarantine facility: “A separate quarantine facility, with the ability to accommodate mammals, birds, 

reptiles, amphibians, and fish should exist. If a specific quarantine facility is not present, then newly 

acquired animals should be isolated from the established collection in such a manner as to prohibit 

physical contact, to prevent disease transmission, and to avoid aerosol and drainage contamination. Such 

separation should be obligatory for primates, small mammals, birds, and reptiles … More stringent local, 

state, or federal regulations take precedence over these recommendations.” 

Quarantine length: “Quarantine for all species should be under the supervision of a veterinarian and 

consist of a minimum of 30 days (unless otherwise directed by the staff veterinarian). Mammals: If during 

the 30-day quarantine period, additional mammals of the same order are introduced into a designated 

quarantine area, the 30-day period begins over again. However, the addition of mammals of a different 

order to those already in quarantine will not have an adverse impact on the originally quarantined 

mammals.” 

Quarantine personnel: “A keeper should be designated to care only for quarantined animals or a keeper 

should attend quarantined animals only after fulfilling responsibilities for resident species. Equipment 

used to feed and clean animals in quarantine should be used only with these animals. If this is not 

possible, then equipment must be cleaned with an appropriate disinfectant (as designated by the 

veterinarian supervising quarantine) before use with post-quarantine animals. Institutions must take 

precautions to minimize the risk of exposure of animal care personnel to zoonotic diseases that may be 

present in newly acquired animals. These precautions should include the use of disinfectant foot baths, 

wearing of appropriate protective clothing and masks in some cases, and minimizing physical exposure in 

some species; e.g., primates, by the use of chemical rather than physical restraint. A tuberculin 

testing/surveillance program must be established for zoo/aquarium employees in order to ensure the 

health of both the employees and the animal collection.” 

Quarantine protocol: “During this period, certain prophylactic measures should be instituted. Individual 

fecal samples or representative samples from large numbers of individuals housed in a limited area (e.g., 

birds of the same species in an aviary or frogs in a terrarium) should be collected at least twice and 

examined for gastrointestinal parasites. Treatment should be prescribed by the attending veterinarian.



Ideally, release from quarantine should be dependent on obtaining two negative fecal results spaced a 

minimum of two weeks apart either initially or after parasiticide treatment. In addition, all animals should 

be evaluated for ectoparasites and treated accordingly. Vaccinations should be updated as appropriate 

for each species. If the animal arrives without a vaccination history, it should be treated as an 

immunologically naive animal and given an appropriate series of vaccinations. Whenever possible, blood 

should be collected and sera banked. Either a -70ºC freezer or a -20ºC freezer that is not frost-free 

should be available to save sera. Such sera could provide an important resource for retrospective disease 

evaluation. The quarantine period also represents an opportunity to, where possible, permanently identify 

all unmarked animals when anesthetized or restrained (e.g., tattoo, ear notch, ear tag, etc.). Also, 

whenever animals are restrained or immobilized, a complete physical, including a dental examination, 

should be performed. Complete medical records should be maintained and available for all animals 

during the quarantine period. Animals that die during quarantine should have a necropsy performed under 

the supervision of a veterinarian and representative tissues submitted for histopathologic examination.” 

Quarantine procedures: The following are recommendations and suggestions for appropriate quarantine 

procedures for small carnivores: 

Required Strongly recommended 

1) Direct and floatation fecal exam 1) CBC/sera profile 

2) Vaccinate as appropriate 2) Urinalysis 

3) Appropriate serology (FIP, FeLV, FIV) 

4) Heartworm testing in appropriate species



Appendix G: Giant Otter Enclosure Design 

Adapted excerpts from the “International Giant Otter Studbook Husbandry and Management Information 

and Guidelines (2005)” for WAZA Website, and with added U.S. units of measurements (Sheila Sykes- 

Gatz 2006). 

The total minimum size enclosure for one giant otter pair should be at least 2,583ft 2 (240m 2 ) and 

when indoor enclosures are needed, they should be a minimum of 807ft 2 (75m 2 ) of the total area. In 

temperate climates, it is necessary that an outdoor enclosure (with or without heated outdoor water) 

provides access to a heated indoor enclosure. Both indoor and outdoor enclosures require the same 

recommended conditions (i.e., land to water ratios, substrates etc.). At least 2 dens (i.e., separable 

rooms) to contain nest boxes/briefly hold animals should be provided, and at least 2-3 nest boxes (or 

natural underground dens) are needed. 

Providing the recommended land to water area ratios (i.e., enough land area), substrate types and 

depths to cover all surfaces, and deep digging area sizes, is just as important as the need to provide a 

swim area in all giant otter enclosures. These are among the most crucial husbandry provisions needed 

to maintain giant otter physical and behavioral health, and they are also necessary to promote successful 

pup-rearing and adjustment to new/unusual situations. To meet these needs, nearly the entire enclosure 

surface area, including dens and nestboxes, must keep sufficiently dry, soft, and sanitary and otters need 

to be able to effectively dig and groom throughout that entire area. The enclosure must also offer a 

sufficient proportion of land, deep digging, and water area. The provisions in the two paragraphs below 

are essential to providing these conditions. 

Different enclosure sizes require different land to water area ratios. As enclosure sizes decrease 

below 2,583ft² (240m²), proportionately increased land area sizes are needed. It is necessary to provide, 

within each indoor and outdoor giant otter enclosure, at least the minimum percentage land area that the 

following formula determines. Convert enclosure sizes to m 2 (ft 2 x 0.093 = m 2 ). For every 1m 2 that the 

(indoor or outdoor) enclosure size is below 240m 2 , multiply that number (without the m² symbol) by 0.1, 

then add the result to the number 60, and this resulting number is the minimum land area percentage (%) 

that the (indoor or outdoor) enclosure requires. For example, a 1,615ft 2 (150m 2 ) enclosure requires a 

minimum of 69% land area, and an 807ft 2 (75m 2 ) enclosure requires at least 76.5% land area. Enclosures 

between 2,583-6,458ft 2 (240-600m 2 ) in size require at least 60% land area. 

It is crucial that nearly the entire area of surfaces/substrates that giant otters are directly exposed to 

are soft, natural, well-draining, not coarse, sufficiently dry and deep, and loose enough so that otters can 

easily dig into them. It is necessary that every indoor and outdoor enclosure surface, including dens, is 

nearly entirely covered with soft pebble-free sand or mulch (i.e., tree bark pieces only), at least 4-8" (10- 

20cm) in depth, or deep soft loose soil with the needed qualities. The substrates used should not have 

gravel, pebbles, rocks/stones



Pools should have deep areas (at least 3.28ft (100cm) deep), shallow areas (which are frequently 

used), and plentiful areas of gently sloping edges for safe pup exits. Varied natural furnishings, e.g., logs, 

tree stumps with roots, cut bamboo, boulders, should be placed on land and in and over pools. Thin logs 

connected with brackets or large sloping rocks placed just behind and bordering pool edges, fence 

covered drains/filters, and drain pipe extensions help prevent substrates from entering water areas, 

cleaning and drainage systems. Furnishings should allow otters, especially pups and parents, easy and 

safe pool access and exits. Enclosure designs, furnishings, and husbandry methods that offer visual and 

acoustic privacy from human disturbances (zoo staff and visitors) during pup-rearing and that allow safe 

gradual introduction of unfamiliar or temporarily separated otters should be provided. Fish should be fed 

exclusively. A variety of good quality fresh water fish, low in thiaminase and fat, should be offered as the 

main diet. Saltwater fish can be offered occasionally.



Appendix H: Description of Nutrients 

Protein: Protein is the main building blocks of animal structure on a fat-free basis. In addition to being an 

important constituent of animal cell walls, protein is one of the nutrients responsible for making enzymes, 

hormones, lipoproteins, and other crucial elements needed for proper bodily functions. Protein also is 

essential for building and repairing body tissue, as well as protecting the animal from harmful bacteria and 

viruses. Furthermore, protein aids in the transportation of nutrients throughout the body and facilitates 

muscle contractions. The requirements for crude protein are effectively requirements for dietary amino 

acids. The requirements are based on the needs of the animal, the quality of the protein, the source of the 

protein, and the digestibility of the protein available. 

Fat: Dietary fat plays an important role in the manufacture of certain hormones. It also plays a crucial role 

in a wide variety of chemical bodily functions. Also, fat functions as a concentrated energy source, serves 

as a carrier for fat-soluble vitamins (Vitamins A, D, E, and K), and provides essential fatty acids. The 

requirements for fat are effectively requirements for dietary fatty acids. 

Vitamin A: Vitamin A is a fat-soluble vitamin essential for maintaining good vision and healthy mucous 

membranes. It contributes to the differentiation and growth of skin tissue and bone formation (including 

teeth), as well as bone remodeling in growing animals, and glycoprotein synthesis. Vitamin A can improve 

skin and hair/fur conditions, help to increase resistance to certain infections, and improve fertility in both 

genders. In many cases, a vitamin A requirement is effectively a requirement for carotenoids (precursors 

to vitamin A). 

Vitamin C (Ascorbic Acid): Vitamin C is a water-soluble antioxidant, which plays an important role in 

biochemical oxidation-reduction reactions, as well as in the formation of collagen, an important protein 

needed for the formation of skin, scar tissue, tendons, ligaments, and blood vessels. Because of this, 

Vitamin C is crucial to an animal’s ability to heal wounds and repair and or maintain cartilage, teeth, and 

bones. It also may reduce infection by increasing immunity. 

Vitamin D: Vitamin D is a fat-soluble vitamin necessary for active calcium absorption, calcium 

metabolism and resorption from bone. Requirements for vitamin D can be totally or partially met by 

exposure to sunlight or artificial UV light (vitamin D is biosynthesized in the skin of animals or in some 

plant cells upon exposure to the appropriate wavelength of UV light; 285-315nm). 

Vitamin E: Vitamin E is a fat-soluble antioxidant that helps to maintain the structure of cellular and 

subcellular membranes by preventing oxidation of unsaturated fatty acids. It also protects tissues from 

free radicals, which are substances known to harm cells, tissues, and organs. Vitamin E is essential in the 

formation of red blood cells and aids the body in Vitamin K utilization. 

Thiamine (B-1): Thiamine is a water-soluble vitamin, which functions as a necessary coenzyme in 

carbohydrate metabolism (converting carbohydrates into energy) and is hypothesized to play a role in 

nerve or neuromuscular impulse transmission. Thiamine also is important in the proper functioning of the 

heart, muscles, and the nervous system. 

Riboflavin (B-2): Riboflavin is a water-soluble vitamin. It functions in two coenzymes: Flavin adenine 

dinucleotide or “FAD” and flavin mononucleotide. Riboflavin is important for growth and the production of 

red blood cells. It also helps the body to release energy from carbohydrates. Microbial synthesis of 

riboflavin occurs in the gastrointestinal tract of some animals, but synthesis appears to be dependent on 

the type of animal and the source of dietary carbohydrate. 

Niacin (Nicotinic Acid): Similar to Riboflavin, niacin is a water-soluble vitamin which functions in two 

coenzymes: Nicotinamide adenine dinucleotide or “NAD” and nicotinamide adenine dinucleotide 

phosphate or “NADP”. Niacin plays a crucial role in assisting the normal functioning of the digestive, skin, 

and nerve systems. Like riboflavin, niacin helps the body to convert energy from food. The niacin 

requirement of many animals theoretically could be satisfied by synthesis of the vitamin from the amino 

acid tryptophan. However, removal rate of an intermediate in the pathway to create niacin is often so 

rapid that virtually none is produced.



Pyridoxine (B-6): Pyridoxine also known as B-6 is a water-soluble vitamin, which aids the body in the 

synthesis of antibodies by the immune system. It also plays a role in the formation of red blood cells and 

helps to promote healthy nerve functions. Pyridoxine is required to produce the chemical activity 

necessary for protein digestion. 

Choline: Choline is an essential nutrient, which contributes to the function of nerve cells. It is a 

component (helps to form phosphatidylcholine, the primary phospholipid of cell membranes) of the 

phospholipid lecithin (found in cells throughout the body) and is critical to normal membrane structure and 

formation. It also functions as a “methyl donor”, but this role can be completely replaced by excess 

amounts of the amino acid methionine in the diet. 

Folacin (Folate, Folic Acid, B-9, Pteroylglutamic Acid): Folacin, or folate, is a water-soluble vitamin, 

which assists the body in the formation of red blood cells. It also plays a major role in the formation of 

genetic material (synthesis of DNA, the hereditary and functioning blueprint of all cells) within all living 

cells. Folacin functions as a coenzyme, which is important at the cellular and subcellular levels in 

decarboxylation, oxidation-reduction, transamination, deamination, phosphorylation, and isomerization 

reactions. Working in conjunction with Vitamin C and B-12, Folacin assists in digestion and protein 

utilization and synthesis. This vitamin may be used to increase appetite and stimulate healthy digestive 

acids. 

Vitamin B-12: Vitamin B-12 is a water-soluble vitamin, which functions as a coenzyme in single carbon 

and carbohydrate metabolism. In addition to playing a role in metabolism, B-12 assists in the formation of 

red blood cells and aids in the maintenance of the central nervous system. 

Pantothenic Acid: Pantothenic acid is a water-soluble vitamin and part of the B vitamin complex. It is 

needed to break down and use (metabolize) food. Pantothenic acid also is needed for the synthesis of 

both hormones and cholesterol. 

Calcium: The mineral calcium (in association with phosphorus) is a major component of the body and is 

largely associated with skeletal formation. It is important in blood clotting, nerve function, acid-base 

balance, enzyme activation, muscle contraction, and eggshell, tooth, and bone formation and 

maintenance. It is one of the most important minerals required for growth, maintenance, and reproduction 

of vertebrates. 

Phosphorus: In addition to acting as a major component of the body and being largely associated with 

skeletal and tooth formation (in conjunction with calcium), phosphorus is involved in almost every aspect 

of metabolism (energy metabolism, muscle contractions, nerve function, metabolite transport, nucleic acid 

structure, and carbohydrate, fat, and amino acid metabolism). Phosphorus is needed to produce ATP, 

which is a molecule the body uses to store energy. Working with the B vitamins, this mineral also assists 

the kidneys in proper functioning and helps to maintain regularity in heartbeat. 

Magnesium: Magnesium is a mineral, which serves several important metabolic functions. It plays a role 

in the production and transport of energy. It also is important for the contraction and relaxation of 

muscles. Magnesium is involved in the synthesis of protein, and it assists in the functioning of certain 

enzymes in the body. 

Potassium: Potassium is a mineral that is involved in both electrical and cellular functions in the body. (In 

the body it is classified as an electrolyte.) It has various roles in metabolism and body functions. 

Potassium assists in the regulation of the acid-base balance and water balance in blood and the body 

tissues. It also assists in protein synthesis from amino acids and in carbohydrate metabolism. Potassium 

is necessary for the building of muscle and for normal body growth, as well as proper functioning of nerve 

cells, in the brain and throughout the body. 

Sodium (salt): Sodium is an element, which the body uses to regulate blood pressure and blood volume. 

Sodium also is critical for the functioning of muscles and nerves. 

Iron: Iron is a trace element and is the main component of hemoglobin (oxygen carrier in the blood), 

myoglobin in muscles (oxygen carrier with a higher affinity for oxygen than hemoglobin), and many 

proteins and enzymes within the body. It also functions in immune defenses against infection.



Zinc: Zinc also is a trace element that is second only to iron in terms of concentration within the body. 

Zinc plays an important role in the proper functioning of the immune system in the body. It is required for 

the enzyme activities necessary for cell division, cell growth, and wound healing. It plays a role in the 

acuity of the senses of smell and taste. Zinc also is involved in the metabolism of carbohydrates. Zinc is 

essential for synthesis of DNA, RNA, and proteins, and it is a component or cofactor of many enzyme 

systems. 

Manganese: Manganese is essential for carbohydrate and lipid metabolism, for synthesis of one of the 

precursors to cartilage formation, and for proper bone formation. Manganese plays a key role in the 

growth and maintenance of tissues and cartilage, specifically proper bone development. It particularly 

aids in development at the ends of bones where new bone formation takes place. This therefore helps to 

reduce the risk of osteoporosis. Manganese also helps to produce certain hormones, metabolizes fat, and 

is part of superoxide dismutase (SOD) an antioxidant. Studies on humans have shown that manganese 

also may lower the frequency of epileptic seizures and enhance immune functioning. 

Copper: Copper is an essential trace mineral present in all body tissues. Copper, along with iron, helps in 

the formation of red blood cells. It also helps in keeping the blood vessels, bones, and nervous and 

immune systems healthy. 

Selenium: Selenium is an essential trace element. It is an integral part of enzymes, which are critical for 

the control of the numerous chemical reactions involved in brain and body functions. Selenium has a 

variety of functions. The main one is its role as an antioxidant in the enzyme selenium-glutathioneperoxidase. 

This enzyme neutralizes hydrogen peroxide, which is produced by some cell processes and 

would otherwise damage cell membranes. Selenium also seems to stimulate antibody formation in 

response to vaccines. It also may provide protection from the toxic effects of heavy metals and other 

substances. Selenium may assist in the synthesis of protein, in growth and development. In humans, 

selenium has been shown to improve the production of sperm and sperm motility. 

Iodine: Iodine is a trace mineral and an essential nutrient. Iodine is essential for the normal metabolism 

of cells. It is a necessary nutrient for the production of thyroid hormones and normal thyroid function. 

US National Library of Medicine

Grams 

1000 

800 

600 

400 

200 

0 

1 11 21 31 41 51 61 71 81 

Day 



Appendix I: Pup Weights of Ex-situ Population Bred Otters 

Asian-small clawed otter (A. cinereus) pup weights (mother-reared) at Institution F (N = 1.3) 

Min weight (g) 

Asha weight (g) 

Tong weight (g) 

Dep weight (g) 

Spotted-necked otter (L. maculicollis) pup weights (mother-reared) at Institution M N=1.0 

Age in days Weight (kg) Age in days Weight (kg) 

28 0.75 98 1.9 

35 0.95 107 1.9 

42 1.2 113 1.95 

49 1.4 120 2.1 

56 1.5 127 2.7(after eating) 

65 1.6 134 2.5 

72 1.65 140 2.6 

77 1.8 148 2.5 

84 1.75 155 2.45 

91 1.9 - -



North American river otter (L. canadensis) pup weights (mother-reared). Data taken from AAZK Zoo Infant 

Development Notebook 1994, Institution N, Institution O, Institution P. 

Males (N = 9) Females (N = 8) 

Age/days Weight (g) Age/days 

Weight 

(kg) 

Age/days Weight (g) Age/days Weight 

(kg) 

1 110-170 32 0.992-1.03 1 170 32 0.971 

2 177-184 33 0.998-1.09 2 177 33 - 

3 193-220 34 1.08-1.11 3 198 34 1.01 

4 204-241 35 1.11-1.14 4 213 35 1.05-1.15 

5 241-276 36 1.13-1.19 5 248 36 1.06 

6 249-298 37 1.16-1.18 6 262 37 1.09-1.23 

7 266-333 38 1.20-1.25 7 298 38 1.13 

8 280-354 39 1.23-1.28 8 333 39 1.15-1.30 

9 325-376 40 1.28-1.34 9 347 40 1.23 

10 353-404 41 1.35-1.36 10 383 41 1.28 

11 364-425 42 1.32-1.41 11 397 42 1.25-1.35 

12 398-453 43 1.35-1.39 12 411 43 1.28 

13 414-475 44 1.40-1.43 13 439 44 1.35 

14 496 45 1.45-1.57 14 454 45 1.39 

15 531-539 46 1.52-1.62 15 489 46 1.43 

16 499-574 47 1.43-1.62 16 517 47 1.34-1.48 

17 595 48 1.59-1.69 17 546 48 1.46-1.60 

18 617-624 49 1.59-1.67 18 560 49 1.58 

19 624-645 50 1.69-1.79 19 609-685 50 1.62 

20 666-680 51 1.62-1.74 20 637 51 1.56 

21 687 52 1.67-1.87 21 652 52 1.53 

22 765-780 53 1.74-1.88 22 660-730 53 1.62 

23 780-808 54 1.74-1.92 23 723 54 1.64 

24 810-843 55 1.71-1.96 24 758-850 55 1.66-1.81 

25 822-858 56 1.54-1.68 25 720-795 56 - 

26 829-872 57 1.71-2.03 26 772 57 1.93 

27 850-872 58 1.87-2.10 27 794 58 1.76 

28 865-910 59 1.90-2.06 28 815-900 59 1.80 

29 907-921 60 1.52-2.12 29 872 60 1.86-1.70 

30 935-978 61 1.97-2.15 30 907 61 1.84-2.33 

31 971-1000 62 1.96-2.24 31 928-1060 62 1.88



Appendix J: List of Commonly Trained Behaviors for Otters 

(Adapted from AAZK Animal Training Committee) 

Commonly Trained Behaviors for: Mustelids, Procyonids, and Viverrids 

Purpose of the list and source of the data: 

The following list of behaviors was derived using data from a 2003 survey conducted by the American 

Association of Zoo Keepers Animal Training Committee (AAZK, ATC). The goal of the survey was to 

census the existence and depth of training programs for species in AZA facilities. For each species 

trained, each respondent was asked to list trained behaviors, types of reinforcement, and conditioned 

reinforcers used. Additional information about facility design, training tools, and general comments was 

also requested. 

Survey results pertaining to the list of behaviors: 

219 AZA facilities were surveyed. There were 71 respondents. 31 of these train species within the 

Mustelid, Procyonid and Viverid taxonomic group. Because many similarities were found within taxonomic 

groups, commonly trained behaviors were compiled to serve as a reference for animal training programs. 

Of the 31 respondents that train within this group, the percentage that train each behavior is listed next to 

the behavior. 

Facility Differences and Individual Animals: 

Not every behavior will work for every animal. The appropriateness of a behavioral goal for an individual 

will depend on management policy and building design of the facility, as well as the needs and disposition 

of the animal. 

The ATC hopes that this data will aid in the design of training programs for the Mustelid, Procyonid, and 

Viverrid taxa. Where appropriate, these commonly trained behaviors can greatly enhance the husbandry 

of species in this group. For questions or comments about this list or the Trained Behaviors Survey, 

please contact the AAZK Animal Training Committee at www.aazk.org. See next page for chart.

Commonly Trained Behaviors for: Mustelids 

MUSTELIDS (Behavior and % of responding institutions) 

Otter (river, small-clawed, & sea) Skunk (striped and spotted) 

Shifting 83% Shifting 33% 

Separations 66% Separations 17% 

Target 79% Target 67% 

Scale 59% Scale 33% 

Squeeze entry/crate 76% Harness training 17% 

Anal/genital present 7% Crate/Squeeze entry 83% 

Back 10% Nail trim 33% 

Belly 21% Back 17% 

Ears 7% Belly 17% 

Otters (continued) Skunk (continued) 

Eyes 14% Paws/feet 33% 

Head presentation 14% Tactile desensitization 50% 

Mouth 21% Station 17% 

Paws/feet 52% 

Sides 10% Badger 

Oral meds 21% Shifting 100% 

Brushing teeth 3% Target 0 

Injection w/ syringe 7% Scale 0 

Stethescope 3% 

In-water behaviors 31% Wolverine 

Vocalization 7% Belly 50% 

Stay (hold) 10% Paws/feet 50% 

Retrivals 17% Separations 100% 

Station 59% Shifting 100% 

Fecal Collection 7% Station 50% 

A to B 10% Squeeze/crate 50% 

Climb 24% Flashlight desensitization 50% 

Flashlight 7% Scale 50% 

X-ray 3% 

Opthalmoscope 3% 

Blood collection 3% 





Appendix K: Enrichment Items Commonly Provided to Otters 

The table below lists items used at various North American facilities for behavioral and environmental enrichment of otters. 

Natural Exhibit Furniture 

Non-edible 

manmade 

Live Food Edibles 

- Soil, sand, mulch - Climbing areas - Boomer balls and - Fish (smelt, - Ice blocks w/fish, 

- Grass, wheat (available in all other products like shiners, goldfish, fish-sicles, fish 

grass, sedges, etc. exhibits, i.e., cliffs, the "spoolie", trout, mackerel, cubes, etc. 

- Trees 

ledges) 

“bobbin” & "ice tilapia salmon)* - krill cubes, clam 

- Vines "vine hoops" - Logs (on land, cube". 

- Crayfish 

cubes, etc. 

- Aquatic plants submerged, - Ice blocks, cubes, - Crickets 

- Frozen or thawed 

- Hay, straw, grass, floating; hollow pops. 

- Giant mealworms sand eels 

leaves, wood and/or solid) - Snow & ice - Earthworms - Fish pieces 

wools as bedding - Rocks (not - PVC cricket feeder - Freshwater clams - Chicken necks 

- Grass piles 

artificial) 

- Buckets 

- Mussels 

- Mice 

- Leaf piles 

- Stream 

- Blankets, burlap, - Krill 

- Whole-fish -frozen 

- Rocks, all sizes for - Sticks 

non-fraying rags, - Eels- naturally or thawed 

play and 

- Browse (leafy towels 

found 

- Whole 

manipulation 

branches on land - Barrels of water - Shrimp 

apples/oranges 

- Knot holes 

and/or floating) - Frisbees 

- Aquatic insects - - Fruit & berries incl. 

- Bark sheets - Slides 

- Tubs of water naturally found grapes, 

- Pine Cones - Tunnels 

- Carpet over board - Mice- naturally blueberries, 

- Mud 

- Stream bed - Rubber-coated found 

strawberries 

- Sod 

- Running water heating pad* - Frogs – naturally - Small pumpkins 

- Bank over-hangs - Holts 

- Astro turf 

found 

and squash 

- Floating wood - Jacuzzi-like jets in - Floating plastic - Grubs 

- Omnivore biscuits 

- Blocks 

pool 

toys 

- Chub 

- Monkey chow 

- Pine needles - Islands in pool - Phone books - Minnows 

- Pigs ears 

- Other animal - Bridges made from - Swim through - Bluegill 

- Frozen blood 

urines 

logs, etc. 

plastic ring - Clams 

blocks, cubes, etc. 

- Powdered scents - Stumps 

- Kids puzzle balls, - Mud minnows - Hard-boiled eggs 

and herbs 

- Natural fiber mat billiard balls, hard 

- Day-old chicks 

- Fresh herbs - Movable sand box balls 

- Crabs 

- Extracts, i.e., - Logs brought from - Pieces of PVC 

- Melons 

vanilla, etc. 

other exhibits pipe and fittings 

- Coconuts 

- Grapevine balls - Log ladder - Kong chews 

- Frozen feline balls 

- Shells 

- Non-sprayed - Metal bowls and 

- Milk bones 

- Turkey feathers evergreen trees pans 

- Screw pine nuts, 

- Corn stalks - Moving soil pots - Plastic tubs and 

unsalted peanuts 

- Blowing bubbles - Hanging logs with bottles 

- Krill patties 

into exhibit 

holes for food - Bread tray 

- Hamster ball w/ 

- Kudzu vines - Snow piles - Plastic slide, 

treat 

- Cow hooves - Piles of ice cubes house 

- Gelatin Jigglers 

- Stock tank 

- Corn on the cob 

- Hanging tub* 

- Yogurt with fish 

- Warm water hose 

- Vari-kennel tubs 

with substrates 

- PVC tube hung for 

climbing in 

- Unsalted ham 

* These items should be monitored for safety.



The following list provides more examples of enrichment initiatives offered to otters at the Institution J and Institution 

L: 

Institution J – ASC otter 

Non-food items 

- Boomer balls & Jolly balls 

- Bowling pins 

- Brushes 

- Bucket lids 

- Beer kegs, feed barrels & trash cans 

- Feed bags 

- Clover clumps 

- Milk crates, Plastic wagons & Plastic logs 

- Water cooler bottles 

- Grass flats/clumps 

- Hang paper maché figures 

- Hollow coconut shells 

- Oscillating fan, wind chimes, & bubble machine (outside 

of enclosure) 

- Large logs, rearrange furniture, etc. 

- Leaves, sand, and rock piles 

- PVC tubes 

- Towels, clothes, blankets 

- Cardboard boxes and tubes (caution needed when 

using paper products that can become wet) 

- Laser pointer 

- Nature tapes 

- Perfume/body sprays & Glad scented sprays 

Institution L – N. A. river otter/ASC otter 

Non-food items 

- Bobbin with smelt rubbed on it 

- Whole coconuts to roll around 

- Yellow pages 

- Bengay ointment inside a boomer ball 

- Log switching between animal exhibits 

- Regular Alka Seltzer ® in PVC tube (very small holes in 

PVC) 

- Corn stalks 

- Blocks of recycled plastic with holes drilled in them to 

dig food items out 

- Crickets in PVC tube feeder 

- PVC shaker toys 

- Milk crates, cardboard box, use with caution 

- Pinecone soaked in scents 

- Extracts – vanilla, almond, lemon & spices 

- Elephant manure 

- Deodorant spray 

- Reindeer antlers & pronghorn sheaths 

- Paper maché 

- Traffic cones 

- Hummus 

- Ice piles 

- Rose petals 

- Burlap sacs 

- Straw piles 

- Reindeer antlers 

- Varied of feeding devices & times 

- Nyla bones 

- Spices and extracts 

- Mirror 

Food items 

- Honey smears 

- Blood popsicles 

- Cooked chicken 

- Crickets 

- Horse meat 

- Meal worms 

- Peanut butter 

- Pinkies 

- Dry cat food 

- Milk bones 

- Tuna 

- Pig ears and cow hooves 

- Painting 

- Mustard or tomato sauce 

- Large black kong toy 

- Floating PVC tube to swim through 

Food items 

- Liver 

- Anchovy paste 

- Hard boiled eggs, apples, pumpkins, 

carrots, blueberries 

- Gelatin jigglers 

- Live crawdads, live trout in pool, crickets 

- Frozen smelt ice blocks 

- Blood popsicles 

- Knuckles 

- Beef hearts 

- Mice and rats



Appendix L: Resources for Enrichment and Training 

Enrichment 

Coe J. 1992. Plan ahead for behavioral enrichment in environmental kaleidoscope: Research, 

management, and design. In: Proceedings of the AAZPA Annual Meeting. Wheeling WV:AAZPA. pp. 

120-23. 

du Bois T. 1992. The Los Angles Zoo environmental enrichment program: We get a lot of help from our 

friends. In: Proceedings of the AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp. 112-19. 

Goss D. 1999. Creating artificial vines for natural zoo displays. AAZK Forum: 26(11):449-51. 

Grams K, Ziegler G. 1995. Enrichment companies and products. AAZK Forum: 22(6):210. 

Hare V, Worley K. 1995. The shape of enrichment: The first generation. In: Proceedings of the AAZPA 

Annual Meeting. Wheeling. WV:AAZPA. pp.180-86. 

Harris A. 1998. Enrichment: Trials, tribulations and magic moments. In: Proceedings of the AAZPA 

Annual Meeting. Wheeling. WV:AAZPA. pp. 271-74. 

Houts L. 1999. Folsom City Zoo pinatas. AAZK Forum: 26(5): 177-78. 

Houts L. 1999. Utilizing young visitors for behavioral enrichment. In: Proceedings of the AAZPA Annual 

Meeting. Wheeling. WV:AAZPA. pp 72-3. 

Houts, L. 2000. Enhancing animal welfare with limited resources. Proceedings of the AZA Annual 

Meeting. Wheeling. WV: AZA. 

Houts L, Greaves J. 2000. Enrichment options- Using Graphics to promote enrichment. AAZK Forum: 

27(7): 307-08. 

Lacinak CT, Turner TN, Kuczaj SA. 1995. When is environmental enrichment most effective? In: 

Proceedings of the AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp. 324-27. 

Lattis, R. 2000. Animal welfare and the AZA. Proceedings of the AZA Annual Meeting. Wheeling. 

WV:AZA. 

Laule G. 1992. Addressing Psychological well being: Training as enrichment. In: Proceedings of the 

AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp. 415-22. 

Laule G, Desmond T. Positive reinforcement training as an enrichment strategy. In: Shepherdson DJ, 

Mellen JD, Hutchins M, editors. Second Nature: Environmental enrichment for captive animals. 

Washington: Smithsonian Institution Press; 1998. pp. 302-313. 

Lindburg D. Enrichment of captive animals through provisioning. In: Shepherdson DJ, Mellen JD, 

Hutchins M, editors. Second Nature: Environmental enrichment for captive animals. Washington: 

Smithsonian Institution Press; 1998. pp. 262-301. 

Markowitz, H. Behavioral enrichment in the zoo. New York: van Nostrand Reinhold Company; 1982. 

Martin S. 1996. Training as enrichment. In: Proceedings of the AAZPA Annual Meeting. Wheeling. WV: 

AAZPA. pp. 139-41. 

McPhee ME, Foster JS, Sevenich M, Saunders CD. 1996. Public perceptions of behavioral enrichment. 

In: Proceedings of the AAZPA Annual Meeting. Wheeling. WV:AAZPA. Pp 248-53. 

Mellen J, Shepherdson D, Hutchins M. Epilogue: The future of environmental enrichment. In: 

Shepherdson, D, Mellen, J, Hutchins, M, editors. Second nature: Environmental enrichment for 

captive animals. Washington, DC: Smithsonian Institution Press; 1998. p. 329-336. 

Mellen J, Sevenich M. 1999. Philosophy of animal enrichment: past, present, and future. In: Proceedings 

of the PAZAAB Conference. Cape Town: South Africa. 

Moore, D. 1997. AZA Minimum Husbandry Guidelines for Mammals: Mustelids. AZA Mammal Standards 

Task Force, 1997. 7pg. 

Moore, D, Reiss, D. 2000. A multi-institutional program addressing welfare, enrichment, husbandry 

training and behavioral research. Proceedings of the AZA Annual Meeting. Wheeling. WV: AZA. 

Myers M. 1999. Feeder designs at the Audubon Park and Zoological Gardens. AAZK Forum: 26(11):369- 

72. 

Seidensticker J, Forthman D. Evolution, ecology, and enrichment: Basic considerations for wild animals in 

zoos. In: Shepherdson DJ, Mellen JD, Hutchins M, editors. Second Nature: Environmental 

enrichment for captive animals. Washington: Smithsonian Institution Press; 1998. pp. 15-29. 

Sevenich MacPhee, M, Mellen, J. 2000a. Framework for planning, documenting, and evaluating 

enrichment programs (and the Director’s, Curator’s, and Keeper’s roles in the Process). Proceedings 

of the AZA Annual Meeting. Wheeling. WV:AZA.



Sevenich MacPhee, M, Mellen, J. 2000b. Training and enrichment models for avian exhibits. Proceedings 

of the AZA Annual Meeting. Wheeling. WV:AZA. 

Sheng S. 1992. The ultimate enriched environment. In: Proceedings of the AAZPA Annual Meeting. 

Wheeling. WV:AAZPA. pp 124-27. 

Shepherdson D. 1992. Environmental enrichment: an overview. In: Proceedings of the AAZPA Annual 


Shepherdson D. 1997. The animal’s perspective: Developing strategies for successful enrichment. In: 

Proceedings of the AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp 486-89. 

Shepherdson, D. Tracing the path of environmental enrichment in zoos. In: Shepherdson, D, Mellen, J, 

Hutchins, M, editors. Second nature: Environmental enrichment for captive animals. Washington, DC: 

Smithsonian Institution Press; 1998. p. 1-14. 

Shepherdson, D, Carlstead, K. 2000. When did you last forget to feed your tiger? Proceedings of the AZA 

Annual Meeting. Wheeling. WV:AZA. 

Shepherdson D, Mellen J. 1993. Environmental enrichment: Carrying forth the message. In: Proceedings 

of the AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp 216-18. 

Shields J. 1995. Behavioral enrichment for the rest of the zoo. In: Proceedings of the AAZPA Annual 


Smith C. 1993. An inexpensive simulated thundershower for small enclosures. AAZK Forum: 20(9):322- 

23. 

Stern S. 1994. Whose life is really being enriched here anyway? Shape of enrichment 3(3). 

Widner K. 1994. Initiation of the zoo-wide enrichment program at the Knoxville Zoo. In: Proceedings of 

the AAZPA Annual Meeting. Wheeling. WV:AAZPA. pp 51-2. 

Williams L. 1996. Ethological considerations for designing behavioral enrichment. Lab Animal. pp 29-33. 

Whorley, K. E. & V. J. Hare. 1995. An overview of the 1995 Environmental Enrichment Conference. The 

Shape of Enrichment. Vol. 4, No. 4, November. 

There are also many enrichment resources available on-line and in print, including: 

"Enrichment Options" – A regular column featuring brief descriptions of ideas published monthly in the 

Animal Keepers' Forum. Published by the American Association of Zoo Keepers, Inc. AAZK 

Administrative Office, Susan Chan, Editor. 3601 S.W. 29th Street, Suite 133 Topeka, KS 66614. 

Phone: (785) 273-9149, Fax: (785) 273-1980. Email: akfeditor@zk.kscoxmail.com. Website: 

www.aazk.org 

"The Shape of Enrichment" Newsletter – A newsletter devoted entirely to enrichment of captive wild 

animals. Published by The Shape of Enrichment, Inc., V. Hare & K. Worley, (eds.). 1650 Minden 

Drive, San Diego, CA 92111. Phone: (619) 270-4273. Fax: (619) 279-4208. E-mail: 

shape@enrichment.org. Website: www.enrichment.org 

The American Association of Zoo Keepers Enrichment Committee: www.aazk.org 

Disney Animal Kingdom - www.animalenrichment.org 

AAZK Enrichment Notebook 3rd ed. 2004 ISBN1-929672-11-X, 

www.aazk.org/2004enrichnotebookcd.php 

Fort Worth Zoo’s Enrichment Online: www.enrichmentonline.org/browse/index.asp 

Training 

Baker A. 1991. Training as a management tool: Creating the climate and maintaining the momentum. In: 

AAZPA Annual Conference Proceedings. Wheeling, W.Va.: American Association of Zoological Parks 

and Aquariums, 563-568. 

Blasko D, Doyle C, Laule G, Lehnhardt J. 1996. Training terms list. Principles of Elephant Management 

School. St. Louis: American Zoo and Aquarium Association, Schools for Zoo and Aquarium 

Personnel. 

Bloomsmith M, Laule G, Thurston R, Alford P. 1992. Using training to moderate chimpanzee aggression. 

In: AAZPA Regional Conference Proceedings. Wheeling, W.Va.: American Association of Zoological 

Parks and Aquariums, 719-722. 

Coe J. 1992. Animal training and facility design – a collaborative approach. In: AAZPA/CAZPA Regional 

Conference Proceedings. Wheeling, W.Va.: American Association of Zoological Parks and 

Aquariums, 411-414.



Coe J. 1992. Animal training and facility design – a collaborative approach. In: AAZPA/CAZPA Regional 


Aquariums, 411-414. 

Dewsbury D. 1978. Comparative animal behavior. New York: McGraw-Hill. 

Grandin T. 1995. Thinking in pictures. New York: Vintage book. 222 p. 

Hediger H. 1950. Wild animals in captivity. London: Butterworths. 207 p. 

Hediger H. 1969. Man and animal in the zoo. London: Routledge and Kegon Paul. 303 p. 

Kazdin A. 1994. Behavior modification in applied settings. Pacific Grove CA: Brooks/Cole Publishing 

Company. 508 p. 

Kleiman D, Allen M, Thompson K, Lumpkin S, editors. 1996. Wild mammals in captivity: Principles and 

techniques. 

Laule G. 1992. Addressing psychological well-being: Training as enrichment. In: AAZPA/CAZPA Annual 



Laule G. 1995. The role of behavioral management in enhancing exhibit design and use. In: AZA 

Regional Conference Proceedings. Wheeling, W.Va.: American Association of Zoological Parks and 


Lyons J. 1991. Lyons on horses: John Lyons’ proven conditioned-response training program. New York: 

Doubleday. 228p. 

Mellen J, Ellis S. 1996. Animal learning and husbandry training. In: Kleiman D, Allen M, Thompson K, 

Lumpkin S, editors. Wild mammals in captivity: Principles and techniques. Chicago: The University of 

Chicago Press, 88-99. 

Mellen J, Sevenich MacPhee M. (2001). Philosophy of environmental enrichment: Past, present, and 

future. Zoo Biology 20: 211-226. 

Pryor K. 1984. Don’t shoot the dog! Simon & Schuster: New York. 

Pryor K. 1995. On behavior. North Bend: Sunshine Books, Inc. 

Ramirez K. 1999. Animal training: Successful animal management through positive reinforcement. 

Chicago: Ken Ramirez and The Shedd Aquarium. 578 p. 

Reichard T, Shellabarger W, Laule G. 1992. Training for husbandry and medical purposes. In: AAZPA 

Annual Conference Proceedings. Wheeling, W.Va.: American Association of Zoological Parks and 


Reynolds G. 1975. A primer of operant conditioning. Palo Alto: Scott, Foresman and Co. 

Sevenich MacPhee M, Mellen J. 2000. Framework for planning, documenting, and evaluating enrichment 

programs (and the director’s, curator’s, and keeper’s roles in the process). In: AAZPA Annual 


Aquariums. 

Shepherdson, D.J. 1998. Tracing the path of environmental enrichment in zoos. In: Shepherdson DJ, 

Mellen JD, Hutchins, M, editors. Second nature: Environmental enrichment for captive animals. 

Washington, DC: Smithsonian Institution Press, 1-12. 

Shepherdson, D., Carlstead, K. 2000. When did you last forget to feed your tiger? In: AAZPA Annual 



Stevens, B., Groff, A., Joseph, S., Mellen, J., Ogden, J., Plasse, R., 2000. Evolving a new species of 

animal care professional: Combining AZA expertise with Disney leadership. In: AAZPA Annual 



Wilkes G. 1994. A behavior sampler. North Bend: Sunshine Books, Inc. 

: 

-S. Maher



Appendix M: Missouri Fish and Wildlife Otter Stomach Contents 

(Missouri Fish and Wildlife (www.mdc.mo.gov/conmag/1999/11/40) MDC Online November 1999, Vol. 60, 

Issue 11. D. Hamilton 

Otter Stomachs containing identifiable fall prey items 

Type Percent 

Crayfish 61 

Fish 51 

Frogs 17 

Muskrats 3 

Ducks 1 

Empty 4 

Ozark otter stomachs containing identifiable fish species 

Species percent 

Bass (sunfish family) 39 

Suckers and Carp 31 

Minnows 14 

Shad 11 

Pike (chain pickerel) 6 

Trout 3 

Catfish 3 

Drum 3 

Unidentified Fish 19 

Age of Game Fish in Ozark Otter Stomachs 

Age Percent 

1-3 years 40 

4-6 years 40 

7-9 years 20



Appendix N: Basic Considerations in the Design and Maintenance of 

Otter Exhibit Life Support Systems 

FOREWORD 

This document fulfills two objectives, 1) providing information on maintaining pools in otter exhibits 

(see Appendices) and 2) providing information to those who are thinking about building a new otter exhibit 

(or renovating an existing one) regarding some of the basic variables to consider. In our experience many 

operational problems stem from the design phase, and it can sometimes be useful to go back to the 

design and construction phase for answers. 

Those with an existing exhibit and/ or with a specific issue may skip past the Planning and Design 

sections straight to the appendices. 

APPENDICES 

1 Glossary of relevant terms 

2 Request for proposal (RFP) [abridged form] 

3 Disinfection 

4 Skimmer boxes 

5 Algae Control 

Those with more detailed questions may contact the author at: juan.sabalones@marylandzoo.org 

INTRODUCTION 

Aquatic systems in zoos include: 

1. Aquariums. 

2. Aquatic exhibits which feature aquatic animals (otters, polar bears, crocodilians, etc.). Fish may 

or may not be displayed. 

3. Water features i.e. ponds, lakes, waterfalls and streams (natural and man made) which may or 

may not feature deliberately introduced exhibit animals. 

4. Interactive experiences such as water rides and children’s “splash zones”. Swimming pool 

regulations may apply. 

5. Various combinations of the aforementioned systems. 

Otter exhibits generally fall into the second category. Anyone planning to build a new otter exhibit or 

to renovate an existing one will have a variety of issues to consider. Each institution will have varying 

circumstances affecting how they approach the project. Nevertheless, there are number of basic steps 

that can be followed regardless of the situation. While it is beyond the scope of this chapter to go into any 

great detail, we have included some appendices to expand on some of the more salient issues. 

PLANNING AND DESIGN 

Typically, a planning and design team will be formed for the initial phase of the project. In addition to 

management and exhibit designers, the expertise of the following parties is needed to achieve the best 

results: 

• Operations 

The husbandry, maintenance, engineering and exhibits, etc. staff have dealt with the 

idiosyncrasies of exhibiting particular flora and fauna. At this initial stage, the designers can use 

their input to help make the system more practical and ergonomic. Once the project gets past a 

certain point in the construction phase, changes becomes very difficult if not, for all practical 

purposes, impossible. In addition, because each aquatic system is a custom installation, problem 

solving is much easier if the operating staff has a good understanding of the design and 

construction phase of their system. 

• Project Manager 

Someone should serve as liaison and coordinator between the staff, the contractors and the part 

of the institution that approves the financial expenditures. Experience in designing, building and 

operating similar exhibits can be very useful. Experience with both roles, i.e. in managing 

contractors and in being a contractor, is also highly desirable. A project manager should strike a 

balance between what is desired and what is possible within the constraints of the overall master



plan/budget and then bring the balance to all relevant parties and work with them to implement it, 

on time and on budget. 

Once the team does its due diligence, if the decision is to hire outside contractors, it is best to have its 

plans as much in order as possible prior to engaging one. This saves on expensive consulting time and 

also helps focus the institution on the task at hand. Regardless of the level of expertise of the contractors 

hired, an institution should maintain this involvement and focus as much as possible throughout the 

project. There is a cliché that you get what you pay for, but the reality is more that many projects could 

have been completed for much less had the planning and design team been more efficient and effective. 

CONTRACTORS 

If they have the wherewithal, they may choose to handle all aspects of such a project in-house, but 

most institutions will have to hire contractors to handle major portions of the job. For new exhibit 

construction or renovation, each institution will have its own procedures for hiring contractors. These can 

generally be categorized as follows: 

A. Hiring a life support designer and a separate installer and managing the operation in- house. 

B. Hiring a design firm (many institutions already have relationships with one) who will then 

subcontract a life support designer and a separate installer. The institution will handle any other 

related work. 

C. Hiring a firm that will handle all of the aspects of building or renovating an entire exhibit (design 

and construction of the exhibit, its décor, life support system and all associated aspects) in what 

is often referred to as a “turn key” operation. 

D. Some combination of these procedures. 

Each institution will have to weigh the positives and negatives of each approach. In terms of what to 

look for in designers and installers, building or renovating an exhibit in a zoo or aquarium represents a 

unique challenge for most contractors, and adapting to that environment takes time and money. The ideal 

contractor will be able to adjust to the particular institution’s environment quickly and successfully: 

• Life Support System/Filtration System Designer Traits 

o Balance: A competent designer will be able to come up with a design that balances the 

needs of the staff with the temporal and budget parameters set by the institution. A bad 

designer can set you back significantly. 

o Practicality: Some designers have the technical capabilities but lack the practical sense 

to translate them into a workable, affordable design in a zoo or aquarium setting. Closely 

scrutinize the consultant’s CV particularly when it comes to previous projects cited. 

Current operators of those projects should be interviewed. Bear in mind other institutions 

may be reluctant to be candid about a contractor’s shortcomings. 

o Experience: The best designers have some operating and maintenance experience with 

the types of systems they are designing and can therefore relate well with the operating 

staff. 

• Installer Traits 

o Expertise: A good installer will be able to take the designer’s plans and build a system 

that works as designed. This is harder than it sounds. 

o Ergonomics: Besides the normal technical competencies one would expect, it is 

important that they demonstrate a good understanding of operational ergonomics (see 

Appendix E) as it relates to such systems. The easier it is for staff to access and operate 

machinery and controls, the more likely they are to properly operate and maintain them. 

Ideally you should inspect their most recent installation. 

o Experience: Installing an aquatic system in a zoo or aquarium is often an unorthodox 

project even for the experienced contractor. Hiring the local swimming pool installer may 

be cheaper, until the time spent by the designer, project manager and operating staff 

guiding them through the project and correcting mistakes is added up. 

WATER QUALITY MEASUREMENT 

“If it cannot be measured, it cannot be managed “ 

Nolan Karras, Speaker of the House, Utah House of Representatives



The key to properly managing any aquatic system is to have the best handle possible on the water 

quality. A good water quality lab is staffed by people with the experience to interpret the data. Along with 

good record keeping, this is an invaluable aid in problem solving. Here is a suggested equipment list: 

• Well Equipped (>$50,000) 

o Liquid Spectrophotometer 

o Ion Chromatograph 

o Travel liquid spectrophotometer 

o Data Sonde 

o Dissolved Oxygen Meter 

o Turbidity Meter 

o Flame Spectrophotometer 

o Temperature compensated pH Probe 

o Refractometer 

o Micro scale 

o Small scale 

o Large scale 

o Micro Pipette 

o Autoclave 

o Large Spinner Plate 

o Small Spinner Plate 

o Testing Kits (CO2, Alkalinity, Hardness) 

o Microbiology Incubator 

o Fume hood 

o Cabinetry / Island 

o Miscellaneous testing materials 

o Miscellaneous safety equipment 

• Moderately Equipped ($20,000-$49,999) 

o Liquid spectrophotometer 

o Colorimeter 

o DO meter 

o Temperature compensated pH probe 

o Refractometer 

o Small Scale 

o Large Scale 

o Small Spinner plate 

o Testing kits 

o Fume Hood 

o Cabinet / storage closet 

o Misc. Testing material / Tools 

o Safety Equipment 

• Minimally Equipped (



After the planning and design team has done their due diligence and developed a plan, they are then 

ready to issue a request for proposal (RFP). 

We have included sections from an actual Request for Proposal (RFP) for the design of a life support 

system for an Otter Exhibit (see Appendix R). Due to its length, it has been abridged but enough remains 

to give you an idea of what is entailed. Other institutions have their own basic version of such a document 

so they will have to modify the pro forma language to suit their requirements. A performance specification 

for the exhibit has been included. The performance of a life support system is measured by its water 

quality and water clarity. Those parameters can and should be quantified as they give the client a point of 

reference with the contractor as to whether the design has met expectations. 

This RFP does not address the issue of temperature control because this particular exhibit did not 

require it. Otters in general seem to be tolerant of a wide range of temperatures depending on the species 

(see Chapter 1.1). Nevertheless, each institution should determine as early as possible if water 

temperature is a concern. If there is any doubt, it should be included in the performance specifications as 

addressing it after the fact is usually very cost prohibitive. 

Part 1: GLOSSARY OF RELEVANT TERMS 

-Juan Sabalones, Instution S 

Aquatic Life Support Systems: There are two types of aquatic life support systems--open and closed. 

Open systems are only possible in certain locations. The primary advantage is (usually) lower cost for 

construction and, potentially, ongoing operations. The primary disadvantage is less control over exhibit 

water quality due to the inability to control the quality of the primary water source. 

Open: An open or flow through system takes water from a source outside the exhibit such as a river, lake, 

stream or even the ocean, and runs it through the exhibit at such a rate as to reduce the need for 

filtration and then returns the water to that source. Such systems are not possible for all locations and are 

far from ideal. The question of how much an open system costs verses a closed one, for example, is 

difficult to answer. It depends upon how open the system is and what the water quality is adjacent to the 

intake. If the otter exhibit is located where an open system is a viable option (i.e. all the proper 

parameters can be met) then the primary considerations, especially in regards to salt water, involve a 

need for some pretreatment (usually some form of sterilization and mechanical filtration). Depending on 

the regulations in your local jurisdiction, there may be a need to treat the water prior to returning it to the 

source. Zoos have long practiced a variation on this known as “dump and fill” where the water leaving the 

exhibit goes straight to a drain. 

Closed: A closed system, ideally, recycles the water so that water changes are performed only because 

of evaporation and should keep all the required water quality parameters at appropriate levels. In reality, 

even the best-designed and best-operated closed systems can only increase the amount of time between 

water changes. Nevertheless, in many locations they may be the only option. 

Closed systems are generally more expensive to build than an open counterpart. This would include 

capabilities such as raw make-up (for salt water systems) and storage, proper mechanical, biological and 

chemical filtration, denitrification, aeration, organics control, and backwash water recovery and especially, 

a top-notch water quality laboratory. 

Water Parameters: The United States Department of Agriculture has certain coliform counts that need to 

be performed for exhibits that fall under the marine mammal category. Otters do not fall under that 

category and therefore water tests are not required by any federal regulatory agency. Nevertheless, it is 

prudent to perform at least weekly water quality tests for bacterial counts and daily tests of chemical 

additive levels. Records should be maintained and available for inspection and reference if problems 

arise. 

Coliform bacteria: Coliform bacterial counts are used to monitor filtration system efficiency and keep track 

of potentially harmful bacteria. Coliform counts should be done at least every other week and more often 

if there are multiple animals using the pool (a policy regarding coliform testing should be set by the 

institution). Often a MPN (Most Probable Number) per 100ml is given as an acceptable limit. However, a 

more accurate measure is the total or fecal coliform count (NOAA 2006). There are no standards 

established for fresh-water otter pools at this writing. At this time, it is suggested that coliform levels be 

maintained at or lower than levels established for rescued pinnipeds by NOAA. These are:



• Total coliform counts should not exceed 500 per ml water, or a MPN of 1000 coliform bacteria per 

100ml water. 

• Fecal coliform count should not exceed 400 per ml. If animal caretakers are routinely exposed to 

pool water, an institution may establish a higher standard of 100 per ml, which is the level 

considered safe for humans; this should be based on institutional policy. 

Temperature: More detailed research is required into optimal water levels for the tropical otter species; 

however, at this time the AZA Otter SSP recommends the following temperature guidelines: 

A. cinereus: The water temperature for A. cinereus should be maintained between 18.3-29.4°C (65-85°F), 

preferably at the warmer end of this scale (Petrini 1998). 

A. capensis, L. canadensis: The water temperature for A. capensis and L. canadensis does not appear to 

be critical. 

L. maculicollis: Water temperature in successful L. maculicollis exhibits has ranged from 8.9-15.6°C (48- 

60°F). Temperatures in the 15.6-21.1°C (60-70°F) range may encourage this species to spend more time 

in the water, however, this is has not been objectively demonstrated at this time. 

P. brasiliensis: Further study into optimal pool temperatures and water temperature exposure 

recommendations for P. brasiliensis is required. Sykes-Gatz (2005) recommends this species should not 

be allowed to swim in unheated water when air temperatures are below 5°C (41°F). Sufficient indoor 

swim areas are needed when seasonal daytime air temperature regularly falls below 15°C (59°F), 

regardless of whether outdoor water is heated. This is particularly true for family groups rearing pups that 

may be held indoors for 4-5 months during cold temperatures. Heating of indoor housing pools is not 

necessary if the ambient air temperature is maintained at recommended levels. 

Each individual institution will have to determine whether their exhibit must be heated or chilled and 

the most efficient way to do that. 

Turnover Rate: The size and capacity of a life support system is based on turnover rate (the given unit 

of time it takes the total amount of the exhibit water to pass through the filtration). This is usually given as 

gallons per minute (GPM). 

The turnover rate is based on the volume of the exhibit combined with the amount of organic matter 

likely to be put into the water by the animals and the environment (leaves and branches, etc.). For 

example, a hippopotamus pool is likely to require a much higher turnover rate than an otter in the exact 

same sized pool. The dirtier the water is, the likelier a higher turnover rate should be employed. 

A proper turnover rate is crucial as a system with a turnover rate that is too low will always be 

laboring to catch up, and a system with a turnover rate that is too high is wasting money. 

Filtration: 

Biological Filtration: This is the use of bacterial colonies to convert ammonia to nitrite and then nitrate 

through nitrification. 

In aquariums with fish, ammonia and nitrite are harmful and should be kept as close to 0.0 as 

possible. With the exception of certain invertebrates in salt water systems, nitrates are not considered 

harmful. Otters should have no problem with ammonia and its derivatives, but an excess of nitrates in an 

otter pool is undesirable as they are a primary source of nutrients for algae. 

Nitrate levels can be kept in check with water changes or by a number of different denitrification 

methods. 

Mechanical Filtration: Mechanical filtration is the mechanical removal of particulate matter suspended in 

the water column before it decomposes. 

The matter is moved to the filter which is then cleansed. Until the filter is backwashed, the particulate 

matter is still in the system. Rapid sand, screen, drum, bag, diatomaceous earth, protein skimmers and 

bead filters are most commonly used in this application. Water clarity is most directly affected by the 

quality of the mechanical filtration. 

Chemical Filtration: The use of any filtering substance designed to chemically attract pollutants in order to 

remove them from the water column. 

• Activated Carbon



• Reverse Osmosis is a method of filtering water by pressing the water against a semi-permeable 

membrane that sits inside filter housing. This membrane allows water molecules to pass through, 

but not others. Minerals, trace and other elements are removed producing pure raw water. 

• Ion Exchange resins are a water filtration medium comprised of natural and synthetic resins in a 

flow-through pouch. They selectively remove ammonia, nitrite, and nitrate from freshwater. 

Part Two: REQUEST FOR PROPOSALS (RFP) [ABRIDGED VERSION] 

OTTER EXHIBIT LIFE SUPPORT SYSTEM RENOVATION. 

The following specifications shall define the scope of this design/build project for installing a Life Support 

System’s (LSS) mentioned above at the Generic Zoo. 

Bidders: 

You are hereby invited to submit a proposal to the Generic Zoological Society, Inc. (the “GZS”) for the 

design and construction of: 

OTTER EXHIBIT LIFE SUPPORT SYSTEM RENOVATION. 

I Bid Documents 

A. RFP Project Description 

B. Project Scope of Work and Specifications attached. 

C. Bid Form (attached to this RFP) 

D. Contractor’s Qualification Form (attached to this RFP) 

E. Generic Zoo, Zoo Contractors Policy 

F. General Condition form 

G. Standard AIA Contract 

H. Prevailing Wage Rates 

PROJECT DESCRIPTION 

Project: Otter Exhibit Life Support System Renovation 

The Otter Exhibit Life Support System Renovation project consists of designing and installing a 

replacement Life Support System for the existing Otter Exhibit as described in the following specifications. 

Contractor Responsibilities: 

1. Where specified standards are in conflict, the more stringent of the two shall apply. 

2. Site verification of specifications during the bid process and post award period. 

3. Visit the site to determine the extent of work, which may or may not be shown in the plans. 

4. Any Owner property damaged as a result of the work associated with this project shall be restored to 

its original condition at Contractor’s expense. Contractor is responsible for documenting existing 

conditions. 

5. Coordinate the marking of any underground utility lines, which are within the proposed limits of 

construction. 

6. The Site shall be kept clean at all times. Contractor shall keep the project site clean from workerrelated 

debris, including surrounding lay-down areas, parking areas, walkways and lawn areas. All 

debris shall be constantly picked up and properly disposed to reduce impact on guest experience and 

so that winds do not carry the debris to other areas of the grounds. 

7. The Zoo has adopted a limited smoking policy. All vendors may only smoke in areas permitted by the 

designated Zoo representatives, and the contractor is responsible for the workmen, including those of 

sub-contractors, disposing of cigarette butts in identified containers. 

Bid Documents: 

1. Request for Proposal 

2. Project Description 

3. Contractor’s Qualification Form 

4. Bid Form 

5. Project Scope of Work and Specifications

6. Generic Zoo Contractors Policy 

7. General Condition form 

8. Standard AIA Contract 



CONTRACTOR’S QUALIFICATION FORM 

The undersigned certifies under oath the truth and correctness of all statements and all answers to 

questions made hereinafter. 

QUALIFICATION OF LIFE SUPPORT SYSTEM CONTRACTOR 

1. In order to be the Life Support System Contractor on this project, bidders are to submit the 

following information to the Zoo not less than ten (10) days prior to the bid opening date. 

2. Experience 

3. Statement certifying that the Contractor has been in business a minimum of Six (6) years and has 

extensive experience in the construction of mechanical life support systems for live animal 

exhibits of similar size and scope. 

4. Contractor must have experience designing and building similar LSS. 

5. References of at least one (1) recent project at an AZA Accredited Zoological Facility within the 

last year, demonstrating experience and ability to install projects of similar size and complexity as 

those described in the RFP. Include the name of the person responsible for the project, phone 

number, and approximate contract amount. 

6. Contractor shall submit full documentation of his/her construction crews and lead personnel, 

detailing the experience of each person listed and their ability to perform all phases of the work to 

the Zoo's satisfaction. 

7. Site Superintendent must have supervised ten (10) or more LSS installations during his/her 

employment with the Contractor 

8. Site Superintendent must have experience in installing ozone systems 

9. Site Superintendent must show proof of 30-hour “OSHA 500” safety certification course. 

10. Contractor shall be a member of the Association of Zoos and Aquariums. 

The Zoo reserves the right to require additional information and/or request a visit to completed work to 

make a determination of the bidder’s qualifications to produce work as described in the Construction 

Drawings and Specifications. 

Project Identification: Life Support Systems Installation 

Name 

Company Name 

Address 

Fax 

Signature 

(If more space is needed, please answer on the back of this sheet) 

Is your organization licensed to do business in the State of……..? 

Title 

Phone 

City/State/Zip 

FEIN 

DATE



Have you ever failed to complete any work awarded to you? If so, note when, where, and why? (If more 

space is needed, please answer on the back of this sheet.) 

Please list all work which your firm will be self performing. 

List Subcontractors. Use back of sheet if necessary. 

List four or more projects executed by your firm within the past three years that were similar in nature and 

scope to this project. Additional projects may be listed on a separate sheet. 

A) Project Name: Location: Year: 

Project Cost: Owner Name and Phone Number: 

B) Project Name: Location: Year: 


C) Project Name: Location: Year: 


D) Project Name: Location: Year: 


If you have State of….. and/or….. MBE/WBE certification for you or any subcontractors, provide 

Certification Numbers, Expiration Dates, and Disciplines/SAIC numbers for which you or they are 

certified. 

Provide names of key personnel to be employed on this project. Indicate the projects listed above 

with which they were involved. 

Name Years Experience Years w/ Firm Projects listed Project Role 

1. 

2. 

3. 

4. 

If you wish, attach photographic documentation of projects listed in above that illustrate work that you 

have completed that is most comparable in style, technique and workmanship to the project.

BID FORM 

Generic Zoological Society, Inc. 

Life Support Systems Installation Request For Proposals 

Deliver To: Construction Department 

Generic Zoo 

Smallville, USA, Planet Earth 



In submitting this bid, the Undersigned declares that they are the only person, or persons, interested 

in said bid, that it is made without any connection with any person making another bid for the same 

contract, that the bid is in all respects fair and without collusion, fraud or mental reservation, and that no 

employee of the OWNER is directly or indirectly interested in said bid, or in the supplies or work in which 

it relates, or in any portion of the profits thereof. 

The Undersigned also declares that they have examined the Request For Proposals, including the 

drawings and specifications contained therein, and that by signing this proposal, they waive all right to 

plead a misunderstanding regarding the same. 

The Undersigned agrees to submit a bid bond, payable to the Generic Zoological Society, Inc., in the 

amount of five percent (5%) of the total bid amount, along with the bid. 

The Undersigned further understands and agrees that they are to furnish all material, equipment and 

supervision to complete entire work for the indicated project and to accept in full compensation therefore 

the stipulated sum or sums as stated herein. 

On acceptance of the proposal for the construction portion of said work, the Undersigned does 

hereby agree to provide the Generic Zoological Society within ten (10) days Payment and Performance 

bonds in the amount of one hundred percent (100%) of the total bid price to provide construction services 

for the consideration named herein. 

The Undersigned agrees to hold open this Bid Proposal for a period of ninety (90) days following the 

submission of this Bid Proposal. 

The Undersigned agrees to provide evidence of insurance coverage along with their bid submission, 

including areas and amounts such as umbrella insurance, general liability, automobile liability, garage 

liability, excess liability, workers’ compensations and employers’ liability. On acceptance of this proposal 

for said work, the Undersigned agrees to provide the Generic Zoological Society, Inc. with a Certificate of 

Insurance adding the Generic Zoological Society, Inc. as an Additional Insured. 

The base bid price of the proposal shall be inclusive of the following: 

• Price to provide the design of the LSS 

• Price to complete the installation of all LSS piping, equipment, accessories, valves in accordance 

with these specifications. 

• A one-year warranty from substantial completion on all parts, labor and material. 

• Performance and payment bonds on the construction phase. 

• Factory representative onsite for initial start-up of all systems. 

• All labor at Davis-Bacon Act Prevailing Wage Rate. 

• Detailed listing of all equipment, labor and materials. 

• Guarantee of performance of LSS to meet criteria. 

• LSS shall be designed and built to maintain the exhibit animals in a healthy condition (see 1.04B). 

• LSS shall be designed and built to maintain high water clarity suitable for overhead viewing. 

OTTER EXHIBIT LIFE SUPPORT SYSTEM RENOVATION 

A. PROJECT SCOPE/SPECIFICATIONS 

1. Contractor 

a. Design and Install a new Life Support System (LSS) for the Otter Exhibit at the 

Generic Zoo. This scope of work shall be split into two contracts. The first phase 

shall cover the design of the LSS. The deliverables for the first phase shall include 

completed designs to meet the project budget and a proposal to provide the



construction of the LSS. The second phase shall include the construction of the 

LSS. 

b. Remove all existing LSS equipment and piping. Please provide an alternate to 

perform this work. 

c. Piping will connect to existing main drains, surface skimmer, returns, waterfall, 

overflow, waste and domestic water lines. 

d. 8 hours LSS Startup and Zoo staff operator training. 

2. Zoo 

a. Exhibits will be emptied of all water and animals. 

b. The Zoo is responsible for the integrity and functioning of existing underground 

and through-wall piping and electrical service. 

c. Provide all electrical connections from existing electrical panels. (See Alternates) 

B. BASE EXHIBIT DATA 

Otter Exhibit 

a. 20,000-gallon fresh water pool. 

b. Exhibit pool is outdoors. 

c. Fresh water provided by existing domestic water supply. 

d. Exhibit maximum capacity to be two river otters. 

e. Exhibit filtration consists of sand filtration with manual backwash operation. 

f. Exhibit disinfection currently done by bromine additions. 

C. LSS DESIGN PARAMETERS 

1. The LSS equipment will be sized to meet or exceed the following minimum requirements. 

If the minimum requirements appear insufficient, provide an explanation for concerns on 

the sizing. Any additions, deletions, or substitutions must be approved by the designated 

representative of the Zoo. 

a. Turnover rate for the Otter system shall be at least 60 minutes. 

b. System shall have flow meters appropriately placed to monitor the turnover rate. 

c. Waterfalls in the exhibits shall be on a separate loop (with a dedicated pump) from 

the LSS. 

d. Both systems shall have skimmers adequately sized and located to deal with the 

heavy leaf litter problem. Skimmer openings shall be screened to protect against 

accidental intrusion by live exhibit tenants. 

e. If sand filters are specified, they will be sized to less than or equal to 10 gpm/sqft 

of filter area at 100% of system flow. Backwash shall be manual or hydraulic in 

operation. 

f. Exhibit disinfection shall be provided by an automated ozone injection treatment 

system. 

g. Ozone system will be sized for a 20% side-stream and a contact time of 2-3 

minutes. It shall include an ozone destruct unit. There shall be no residual ozone in 

the exhibit. 

h. There shall be a trickle filter sized for the full system flow at 15 gpm / sq. ft (of 

tower cross section). The tower shall have enough height to provide adequate gas 

exchange and prevent air entrainment in the exhibit. 

i. All domestic freshwater supply lines shall have flow meters. 

j. All equipment shall be able to be disconnected with unions or flanges from the 

system. 

k. All equipment will be fully bypassable. 

l. Valved water sampling ports will be provided in the LSS equipment area. 

m. Two primary filtration pumps that will each have the capacity to run the LSS at 

design capacity will be installed so that a full backup pump is accessible with the 

turn of two valves. Piping on the suction side of the filter pumps shall be sized to 

handle the designed system flow at less than 4 ft/second. Piping on the discharge 

side of the filter pumps shall be sized to handle the designed system flow at less



than 6 ft/second. Gravity piping will be sized to accommodate the design flow at 

less than 2.5 ft/sec. 

n. Contractor to verify new equipment will fit in the area of the existing pump room 

obtained by removing existing equipment. 

o. Contractor to verify prior to bid that existing piping to remain is appropriately sized 

for new LSS. 

p. Contractor to install all systems according to local code and arrange for all 

inspections. 

2. The LSS contractor will be required to guarantee the performance of the LSS once 

installed. If in the opinion of the bidding contractor, the above criteria will not meet the 

quality standards set forth here or meet the LSS contractor’s own standards, those 

deficiencies should be detailed in the proposal. The design must allow the system to 

achieve the following parameters under normal operating conditions : 

Parameters 

Temperature (appropriate to species specific range) 

Calcium Hardness 100- 200 

ORP 300-400 pool, 750 - 800 contactor 

Ozone in Water 0 ppm 

Total Alkalinity 80-120 ppm 

Dissolved Oxygen 90-100% saturation 

Free ammonia 0 

Nitrite (NO2)



1) The contractor shall install all piping, valves, and equipment in a manner 

and in locations to avoid obstructions and keep openings clear. Pipe and/or 

equipment shall not be installed where it will present a potential tripping 

hazard or below 7’-0” above finished floor, where it would be a potential 

head knocking hazard. Installation shall permit direct access to all valves 

and pieces of equipment that will require maintenance. The contractor shall 

make any changes as directed by the owner, at no additional expense, 

which may be necessary in order to accomplish this purpose. 

2) Before being placed in position, all pipe, pipe fittings and accessories shall 

be cleaned, and shall be maintained in a clean condition. Piping shall be 

installed and aligned in accordance with the Drawings with a tolerance of + 

1/8-inch in the horizontal and vertical directions. 

3) All work specified and not clearly defined by the Drawings shall be installed 

and arranged as directed and in a manner satisfactory to the owner. 

b. Installation of Pressure Filter Gravel and Sand 

1) Preparation of Filters: Before placing the media, the contractor shall 

determine that all holes in the underdrain system are open. 

2) Placing of Gravel: The gravel shall be deposited in such a manner as to 

avoid endangering the header laterals. Each layer shall be brought up to the 

required elevation and made level over the entire filter bed area, and shall 

be smoothed down to a true surface. Care shall be taken not to injure 

equipment piping and coatings in the filter units by walking on or dropping 

gravel upon them. 

3) Placing of Sand: In placing of the sand in the filters, extreme care shall be 

taken to avoid disturbing the internal piping. Extreme care shall be taken to 

protect the internal filter coating. Sand shall be placed to the appropriate 

depth specified and shall be level. 

c. Installation of Piping 

1) All PVC Pipe will be transported, stored and installed with regard to 

manufacturer's recommendations. Bolting of PVC flanges shall be in 

accordance with manufacturer's recommendations and shall not be unduly 

stressed through the use of excessive torque while tightening bolts. Use of 

torque wrench will be required. 

2) All life support piping shall be flushed clean prior to connection to equipment 

or tanks. 

d. Installation of Pumps and Motors 

1) Pumps and motors shall be installed in accordance with the Manufacturer's 

Recommendations. 

2) Pumps shall be installed with an isolation valve on the suction side of the 

pump. This valve shall be a true-union ball valve unless it is 4” or greater, in 

which case it will be a gate valve. A compound gauge shall be installed 

between the pump and the isolation valve. 

3) Pump shall be installed with an isolation valve on the discharge side of the 

pump. This valve shall be a butterfly valve. Between the pump and the 

isolation valve, a swing check valve is required. A pressure gauge shall be 

installed between the check valve and the isolation valve. 

4) Unless otherwise noted, all pumps shall be installed on concrete 

housekeeping pads. 

e. System Startup 

1) The contractor shall provide one full-time mechanical technician who is 

familiar with the system for a period of 3 days to aid the owner in starting 

and operating the system.

PART 3: DISINFECTION 



Bromine: Bromine is unsuitable for outdoor usage as it is broken down very quickly by sunlight 

Ozone-Chlorine comparison: The common perception is that ozone systems are more elaborate and 

expensive than chlorine. In truth, both systems will require that water parameters be measured. If the 

operator starts with more or less a blank slate, the level of sophistication required to safely and effectively 

operate and maintain a properly designed chlorine disinfection system vs. the level of sophistication 

required to safely and effectively operate a properly designed ozone disinfection system may just about 

be equal. 

Indeed, one could argue that the chlorine system operator needs a higher level of sophistication and 

dedication and needs to work harder and be more attentive than if they had an ozone system. As we are 

talking about otters, when comparing an ozone system to a chlorine system, the latter system is more 

likely to harm the animals (or their human caretakers). 

Ozone: If you have a well designed, installed and maintained system, the operator will keep the various 

filters and probes clean. They might occasionally turn a few dials but other than that, operation consists 

primarily of monitoring the system and recording data. Every year or so you should get a visit from an 

ozone tech for the more serious maintenance issues depending on your system, but that is about it. 

Advances in design now allow us to use much smaller dosages of ozone than in the past with the 

resulting decrease in safety concerns. 

The oxidation process with ozone occurs in the contact chamber away from the animals, and any 

residual ozone is unlikely to be anywhere near as harmful, if at all, to the otters as chlorine residuals. The 

by-products of ozonation can be dealt with. 

Chlorine: In the United States, many municipalities add chlorine to their water, and readings from tap 

water of 1ppm or higher are possible. For this reason, many aquariums run their city supplied water 

through carbon filters prior to use. While otters generally show no adverse effects from these levels, it is 

not known what the overall impact is to their health and the water repellency of their coats. For this 

reason, the AZA Otter SSP recommends that otters should not be exposed to chlorine levels higher than 

0.5ppm for prolonged periods, and ideally, chlorine should be kept at a non-detectable level. The addition 

of sodium thiosulfate will neutralize any residual chlorine. The most efficient and effective way to do this is 

to use an automated system (www.polarispool.com/products/details.asp?ID=34). 

This would cost you about $5000.00 to buy and install. Because chlorine is most effective between 

the pH of 7.2-7.8, such automated systems draw from sodium hypochlorite barrels and muriatic acid 

barrels. Without an automated system, the pool will have to be tested daily and chemical adjustments 

made manually. Even with the automatic controller, this system is inherently more labor intensive than an 

ozone system. 

The oxidation process with chlorine occurs in the water with the animals that are exposed directly to 

the chemicals. Therefore an overdose of chlorine or muriatic acid would affect the animals. Increasingly, 

evidence is mounting that chlorinated pools may cause health problems in humans 

(www.swimming.about.com/od/allergyandasthma/a/cl_pool_problem_3). 

Therefore all things being equal, one has to wonder how sensible and ethical it is to continue using 

chlorine with otters. Chlorination produces carcinogenic byproducts know as Trihalomethanes (THM) 

(www.epa.gov/enviro/html/icr/gloss_dbp). Sooner or later, this may become a regulatory issue and we 

have no way of removing THM from the process. 

In the end, the decision on the type of disinfection to use is an equation. Your particular institution 

may have a set of circumstances (access to highly trained personnel and sophisticated water quality 

measuring equipment, inexpensive sources of sodium hypochlorite and muriatic acid, lots of cheap labor, 

etc.) that will sway the equation towards chlorine. And in cases where algae is out of control, chlorine may 

be the most immediate and cost effective solution. 

However, all things being equal, our experience has been that when you add up the total cost of 

doing it correctly over a number of years, having an ozone disinfection system works out to be cheaper, 

less dangerous, and far less labor intensive. Time is money.



Basic Manual Chlorination Device: In manually applied situations, the two most common forms of 

chlorine used are granular and liquid. 

Granular: Granulated chlorine is designed to dissolve slowly over time so it needs someplace (typically a 

skimmer or a floating chlorinator) where it can dissolve away from direct human or animal contact. 

DERBY DUCK FLOATING CHLORINATOR 

Large capacity holds up to six 3” tabs (or 1” tabs or sticks). 

Adjustable to regulate the amount of chlorine dispensed. Made of 

heavy duty resin – child resistant lock. 

Liquid: In its liquid form (sodium hypochlorite) the chlorine is more concentrated. It would be used if the 

need is to raise the levels of chlorine quickly. Sodium hypochlorite is very corrosive and proper 

procedures (and suitable personal protection) need to be taken when using it. 

Gas: Chlorine gas is also commercially available, but this is typically for industrial applications and should 

be considered too involved and dangerous for most institutions. 

Automated Chlorination Device: The Polaris Watermatic Pro System L-1 and L-2SC are complete 

systems for feeding sodium hypochlorite (liquid) and muriatic acid (liquid). Easy to install, operate and 

maintain, these systems include controllers that plug into any 120V outlet and provide built-in receptacles 

for the peristaltic pumps. The L-2SC System further provides easy-to-use graphic displays and 

adjustable-rate pumps. 

Benefits: 

- Complete liquid system 

- Integrated Flow Cell 

- Easy to install and maintain



Drawbacks: 

- Need to buy, handle and store toxic chemicals such as sodium hypochlorite and muriatic acid. 

- Exposes animals and workers directly to potentially caustic substances. 

Chlorine Removal: Chlorine can be used, if necessary, when the otters are not present. Chlorine is a 

volatile chemical and sufficient aeration can lower levels dramatically in twenty four hours. In situations 

where time is an issue, sodium thiosulfate can be added and run through the system for an hour before 

the otters are allowed access again. To remove chlorine with sodium thiosulfate: 

Amount Sodium Thiosulfate (grams) = aquarium volume (gal) x 0.0038 x [7 x tested chlorine level 

(mg/l)] 

Example: 

Problem: 20,000 gallon tank. Chlorine test results show 0.5mg/l of chlorine in the system following water 

change and refill. To remove the chlorine you will need ____ grams of sodium thiosulfate. 

Solution: 20,000 x 0.0038 x (7 x 0.5) = 266 grams Sodium Thiosulfate

Part 4: SKIMMER BOXES 



All skimmer boxes should be protected with appropriate screening to prevent live animals from getting 

trapped, though this should not be a concern with healthy animals. Screening also prevents foreign 

objects from harming pumps. A skimmer box can be placed outside the exhibit tank or inside 

There are commercially available skimmers typically found in pond and swimming pool supply 

houses, but they are usually too small for most large otter exhibits so chances are skimmer boxes must 

be fabricated by the installer. Ideally, the pool should have outflows at three points: 

A drain from its lowest point 

A midlevel drain 

A skimmer box drain 

Skimmers should be mandatory, especially in outdoor pools. They aid in the removal of floating 

organic material (leaves, twigs, etc.) most commonly associated with such pools.

SS 









Part 5: ALGAE CONTROL 

The presence of algae in an otter exhibit pool has some benefits. A manageable amount can help 

control nitrate levels. However, if the level of algae exceeds manageable limits, it makes the water 

unsightly, reduces oxygen levels, and potentially reduces water circulation. 

Algae need light, appropriate temperature, and nutrients. Control is a concern, particularly in outdoor 

otter pools exposed to significant amounts of sunlight. By providing for a suitable amount of shade and 

controlling the amount of available nutrients, it is possible to keep algae manageable. Therefore 

controlling algae is best addressed in the design phase of the exhibit when solutions to these concerns 

can be built into the exhibit design. Some of them can be added after the fact but it is usually more 

expensive and troublesome to do so. 

Light: If the exhibit is located where the amount of direct sunlight is a problem, a suitable shade structure 

should be found or fabricated. Coloring agents added to the water can limit the penetration of light to the 

algae growing below the water's surface by coloring the water. It seems most of them turn the water 

various shades of blue. Without light, the algae cannot grow. If you have Ultra Violet (UV) disinfection 

devices on your filtration system, coloring agents will render them useless and the colored water may be 

an aesthetic issue. They claim to have no effects on animals, but blue tinted otters would be very 

unsettling. 

Trees provide shade but they may also provide falling leaves and branches which foul the water. 

They also may be toxic in some instances. In such cases, it would be worthwhile to mount some sort of 

structure, such as a net, to deflect the material from the water. 

Nutrients: Algae need nutrients to grow. Nitrates and phosphates are the main concerns. Nitrates come 

mostly from decaying organic material, found in a variety of sources such as animal waste, 

uneaten/leftover food, or decaying leaves and branches. Good hygiene and a properly designed filtration 

system will help. In an outdoor exhibit, a raised lip around the pool will divert rain runoff around the pond 

and help keep dirt out. 

Phosphates usually come in the source water or in food for the animal. At this writing, they can be 

dealt with by filtering the source water to remove phosphates (expensive) or by treating the system with 

lanthanum chloride in its various commercial forms. To the best of my knowledge, lanthanum chloride is 

not harmful to otters but you should check with your vet regardless. 

If possible, additional floating plants in the exhibit can compete with the algae for nutrients and also 

reduce the amount of available light. 

Mechanical Filtration and UV Sterilizers: Mechanical filtration moves undesirable organic matter from 

the water column to the filters. This means the algae have a reduced amount of nutrients available to 

them. The organic material is typically retained in the filtering material, so keeping the filter regularly 

backwashed and working efficiently is vital. 

UV sterilizers are installed in line after the mechanical filter. The cleaner and clearer water helps UV 

sterilizers function properly. The clearer the water, the more effective the UV. When water is pumped 

through the UV sterilizer, the ultraviolet light that is emitted will break down the cell wall of the algae and 

the algae will then die. UV’s also have disinfectant capabilities. 

UV dose = UV intensity X exposure time. 

Make sure to get the appropriate sized UV sterilizer for your pond, and also make sure that the water 

flow through the UV allows for sufficient exposure time. Note that the effectiveness of UV drops 

dramatically as the temperature goes down. Of course, in terms of algae control in outdoor exhibits, algae 

populations are likely to drop down with the temperature. 

Water Additives: Most water additives developed for algae control come from the swimming pool and 

ornamental pond industry. Many of these products are toxic, not only to algae, but to other forms of life. It 

is important to take this into account when considering them for use in an otter exhibit. Always check the 

label and the MSDS sheet to make sure that it is safe for the life in your pool. 

Additives can either be algiscidal or algistatic. Algiscidal products kill the algae outright. Most metal 

ion complexes (copper, silver) fit into this category. Other products, such as barley straw, suppress the 

growth of algae in some fashion and are usually used in a preventative manner. It is important to 

understand at what stage the algae problem is so that the correct product is used. For example, barley 

straw, an algistatic substance, is not as effective after the algae in your exhibit have bloomed. At that



stage, it might be best to drain the pool, scrub the algae off the surfaces, refill the pool and then use the 

algistatic substances to help hold down a recurrence of the problem. 

The use of bags of barley straw to filter out algae has been quite publicized. The following links to 

publications by Carole A. Lembi, Professor of Botany at Purdue University, can provide greater insight 

into Aquatic Plant Management and the use of barley straw in algae control. 

www.ces.purdue.edu/extmedia/WS/WS_21.pdf 

www.btny.purdue.edu/pubs/APM/APM-1-W.pdf 

References and Further Reading: 

Dawes, J. The Pond Owner's Problem Solver. Tetra Press. Blacksburg, Virginia; 1999. 

May, PJ. The Perfect Pond Detective Book 1. Kingdom Books. Waterlooville, England; 1998.



Appendix O: Female Otter Reintroduction Plan 

This plan was developed to reintroduce two females separated for several weeks. It is offered as a 

template to follow when introducing females or any unfamiliar animals. Introductions should be planned in 

advance and based on individual institution policies, physical exhibit design, and individual animals. 

• Begin with one holding cage between them and visual access through mesh in holding. 

• Facility allowed for each animal to have one side of a separated exhibit. They were still with one 

holding cage between them at night. 

• After a few days, the otters were given continual access to side by side holding dens throughout 

the day and night. 

• Otters were switched between exhibit sides every three days to reduce the chance of creating a 

territory. 

• Historically, otters have been introduced in holding then allowed on exhibit. 

• The first day of introductions seems to the have the most signs of aggression. 

• Start with short periods of introduction in holding, varying the time between 20 – 60 minutes. The 

time will vary according to signs of aggression. 

• Once the otters appear to be more comfortable with each other progress to introducing them in 

holding twice a day for a period of 30-60 minutes. 

• When both introduction sessions start going well, allow them access to the exhibit. 

• Watch for signs of aggression and adjust time together accordingly. 

• Each otter should be separated with visual access to one another overnight until confident that 

there is no chance for injury. 

Positive behaviors to look for during the introduction are playful wrestling, muzzle touching, social 

grooming, face pawing, submissive rolling over, resting together, and friendly vocalizations. These 

vocalizations are chirping, grunts or chuckling. 

Aggressive vocalizations are screaming, snarling, growling or grunting. Signs to look for requiring 

separation include aggressive chasing, aggressive wrestling, tension while dominance mounting, fighting 

with a lot of screaming, fighting with injuries, or sign of one trying to drown the other in the pool. 

Tools recommended having on hand when beginning introductions: 

• 3 brooms 

• 4 pairs of gloves 

• 2 hoses that are ready 

• 2 fire extinguishers 

• 3 mammal nets 

• 1 air horn 

• Tongs 

• Snake hook 

• Extra fish and treats 

By: Jessica Foti 

Date: June 4, 2006

5 

Matrix is still in development, photos of each body condition are being sought. 

Photo/Drawing 

Appendix P: Otter Body Condition Matrix 

Cheryl Dikeman, NAG Advisor 

SCORE 1 Emaciated 2 Poor 3 Ideal 4 Solid 5 Obese 

General Condition 

Neck and Shoulders 

Abdomen and Waist 

Hindquarter 

Vertebrae and Rib Cage 

No obvious fat and loss of 

muscle mass. Lumbar 

vertebrae all visible, ribs 

visible, obvious abdominal 

tuck. Ilia wings pronounced. 

Poor coat. 

Pronounced scapula and 

lack of muscle over the 

shoulders, Noticeable 

shoulder skeletal region. 

Very pronounced waist and 

severe abdominal tuck 

Pronounced and very 

obvious hip and iliac region. 

All vertebrae visible. Visible 

ribs. 

Lean, minimal muscle 

mass 

Visible scapula with little 

muscle over the 

shoulders, thin neck. 

Visible delineation behind 

shoulders. 

Visible waist behind the 

ribs. No visible abdominal 

fat present. 

Pelvic bones visible. 

Tops of lumbar and 

thoracic vertebrae and 

ribs slightly visible and 

definitely palpable. 

Optimum body fat and muscle 

tone, well proportioned, ideal coat 

condition. 

Smooth lines over shoulders and 

scapula. Slight delineation behind 

shoulder region. 

No visible abdominal tuck. Some 

distinguishable abdominal fat 

present; however, not obvious. 

Hips and pelvis slightly visible and 

palpable but not obvious. 

Smooth lines over topline and 

throughout body. No visible ribs or 

vertebrae. 

Noticeable fat deposits 

throughout body. 

Smooth lines over shoulders 

and scapula. No delineation 

behind shoulder region. 

Some rounding in the 

abdominal region. Noticeable 

abdominal fat. Waist is not 

visible. 

No skeletal visibility in 

hindquarter. Smooth lines 

over entire quarter. 

Some fat evident over 

vertebral bodies and/or ribs 

Obvious fatty deposits, no 

definition between shoulder, 

stomach and pelvic regions 

No definition, very thickened 

neck region. Obvious fat 

deposits over top of shoulders 

and in neck region. 

Obvious abdominal fat 

deposits and large protruding 

waist region. Abdominal fat 

pad drops below the rib cage. 

Fat deposits obvious over hind 

limbs. Fat pad obvious on 

tailhead. 

Extreme fat pad over rib cage 

region. Heavy fat deposits over 

vertebrae. 

154

Otter (Lutrinae) Care Manual - Association of Zoos and Aquariums

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